Geological Carbon Sequestration and its Influence on Subsurface Microbial Diversity and Metabolic Carbon Cycling
Mariana Erasmus
Geological Carbon Sequestration and its
Influence on Subsurface Microbial Diversity and
Metabolic Carbon Cycling
By
Mariana Erasmus
Submitted in fulfillment of the requirements for the degree
PHILOSOPHIAE DOCTOR
In the
Department of Microbial, Biochemical and Food Biotechnology
Faculty of Natural and Agricultural Sciences
University of the Free State Bloemfontein
Republic of South Africa
August 2015
Promotor: Prof. E. van Heerden
Co-promotors: Prof. D. Litthauer : Prof. T.C. Onstott : Prof. T.J. Phelps : Dr. T. Surridge
I dedicate this thesis to my godchild, Hayley Shardelow.
“Words of wisdom are spoken by children at least as often as
scientists.”
~ James Newman
“The more I study science, the more I believe in God.”
~ Albert Einstein
“If at first, the idea is not absurd, then there is no hope for it.”
~ Albert Einstein
“The value of a college education is not the learning of many facts, but
the training of the mind to think.”
~ Albert Einstein
“It isn't what you learn that makes you successful, but what you use of
what you learn.”
~ Percy H Whiting
CONTENTS
CONTENTS ...... i
ACKNOWLEDGEMENTS ...... vii
LIST OF SYMBOLS AND ABBREVIATIONS ...... x
LIST OF FIGURES ...... xvi
LIST OF TABLES ...... xxviii
CHAPTER 1: LITERATURE REVIEW ...... 1 1.1 GLOBAL CLIMATE CHANGE AND THE GREENHOUSE EFFECT ...... 1 1.2 CARBON SEQUESTRATION ...... 5 1.2.1 Geological Sequestration ...... 6 1.2.2 Ocean Sequestration ...... 8 1.2.3 Terrestrial Sequestration ...... 9 1.2.4 Mineral Sequestration ...... 10 1.3 CARBON SEQUESTRATION IN SOUTH AFRICA ...... 10 1.4 THE CARBON CYCLE ...... 12 1.5 THE DEEP SUBSURFACE AND CARBON CYCLING ...... 13 1.6 CONCLUSIONS ...... 17 1.7 REFERENCES...... 19
CHAPTER 2: INTRODUCTION TO PRESENT STUDY ...... 28 2.1 ABSTRACT ...... 28 2.2 POSSIBLE OCCURENCES DURING CARBON SEQUESTRATION ...... 29 2.3 REFERENCES...... 31
Contents i
CHAPTER 3 - SAMPLING, CHARACTERIZATION AND MICROBIAL DIVERSITY OF THE SITE DESIGNATED TO MIMIC CARBON SEQUESTRATION CONDITIONS ...... 35 3.1 INTRODUCTION ...... 35 3.1.1 Geological Sequestration Sedimentary Rock ...... 37 3.2 AIMS OF THIS CHAPTER ...... 38 3.3 MATERIALS AND METHODS ...... 39 3.3.1 Sampling Requirements ...... 39 3.3.2 The Study Site ...... 40 3.3.3 The Borehole ...... 41 3.3.4 Fissure Water and Gas Sampling and Analyses ...... 42 3.3.4.1 On-Site Physicochemical Analyses ...... 43 3.3.4.2 Collection of Fissure Water for Geochemical Analyses ...... 45 3.3.4.3 Collection of Gas and Fissure Water for Gas Chemistry and Isotope Analyses ...... 47 3.3.5 Sandstone Sampling and Analyses...... 50 3.3.5.1 Crushing and Sterilizing ...... 50 3.3.5.2 X-Ray Fluorescence Analyses ...... 51 3.3.6 Diversity Analyses of the Deep Subsurface Microbial Biome ...... 51 3.3.6.1 Cornelius® Canister Sampling ...... 52 3.3.6.2 SterivexTM Filter Sampling ...... 54 3.3.6.3 Tangential Flow Filtration ...... 54 3.3.6.4 Massive Filter Sampling ...... 55 3.3.6.5 Polyvinyl Chloride Cartridge Sampling ...... 58 3.3.6.6 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis ...... 59 3.3.6.6.1 Genomic DNA Isolation ...... 59 3.3.6.6.2 Polymerase Chain Reaction ...... 59 3.3.6.6.3 Nested Polymerase Chain Reaction...... 61 3.3.6.6.4 Denaturing Gradient Gel Electrophoresis ...... 62 3.3.6.6.5 Gel Extraction, Re-amplification and Purification ...... 62 3.3.6.6.6 Sanger Sequencing and Analyses ...... 63 3.3.6.7 Microbial Diversity Assessments using Targeted 16S and 18S rRNA Gene Sequencing ...... 65 3.3.6.7.1 Genomic DNA Isolation ...... 65 3.3.6.7.2 MiSeq Illumina® Sequencing and Analyses ...... 66
Contents ii
3.3.6.7.3 Roche GS Junior Sequencing and Analyses ...... 66 3.3.6.7.4 Ion Torrent PGMTM Sequencing and Analyses ...... 67 3.3.6.8 Microbial Diversity Assessments of the Fissure Water using Visual Methods ...... 67 3.3.6.8.1 Negative Stain ...... 68 3.3.6.8.2 Live/Dead Stain ...... 68 3.3.6.8.3 Gram Stain...... 69 3.3.6.8.4 Scanning Electron Microscopy ...... 69 3.3.7 Summary of all Samplings and Collaborators ...... 71 3.4 RESULTS AND DISCUSSIONS ...... 73 3.4.1 Fissure Water and Gas Analyses ...... 73 3.4.1.1 On-Site Physicochemical Analyses ...... 73 3.4.1.2 Geochemical Analyses ...... 74 3.4.1.3 Gas Chemistry and Isotope Analyses ...... 76 3.4.2 Sandstone X-Ray Fluorescence Analyses ...... 77 3.4.3 Characterization and Diversity of the Deep Subsurface Microbial Biome .. 78 3.4.3.1 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis ...... 78 3.4.3.1.1 Genomic DNA Isolation and Polymerase Chain Reaction ...... 78 3.4.3.1.2 Nested Polymerase Chain Reaction...... 79 3.4.3.1.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses ...... 80 3.4.3.2 Microbial Diversity Assessments using Targeted 16S and 18S rRNA Gene Sequencing ...... 82 3.4.3.3 Microbial Diversity Assessments of the Fissure Water using Visual Methods ...... 88 3.4.3.3.1 Staining ...... 88 3.4.3.3.2 Scanning Electron Microscopy ...... 92 3.5 CONCLUSIONS ...... 94 3.6 REFERENCES...... 96
CHAPTER 4: MONITORING OF GROWTH AND LOW PRESSURE STUDIES ...... 109 4.1 INTRODUCTION ...... 109 4.1.1 Eubacterium limosum ...... 109 4.2 AIMS OF THIS CHAPTER ...... 110
Contents iii
4.3 MATERIALS AND METHODS ...... 111 4.3.1 Medium Preparation for the Positive Control Microorganism ...... 112 4.3.2 Identification of the Positive Control Microorganism ...... 112 4.3.2.1 Genomic DNA Isolation and Polymerase Chain Reaction ...... 113 4.3.2.2 Ligation of the 16S rRNA Gene ...... 113 4.3.2.3 Transformation of the 16S rRNA Gene ...... 114 4.3.2.4 Analyses of the 16S rRNA Gene ...... 115 4.3.3 Cultivation Conditions for Eubacterium limosum ...... 115 4.3.4 Cultivation Conditions for the Subsurface Biome ...... 118 4.3.5 Microbial Diversity Assessments of Eubacterium limosum and the Subsurface Biome using Visual Methods ...... 119 4.3.6 Microbial Diversity Assessments of the Subsurface Biome using Denaturing Gradient Gel Electrophoresis ...... 120 4.3.6.1 Genomic DNA Isolation and Polymerase Chain Reaction ...... 120 4.3.6.2 Nested Polymerase Chain Reaction ...... 120 4.3.6.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses ...... 121 4.4 RESULTS AND DISCUSSIONS ...... 122 4.4.1 Molecular Identification of the Positive Control Microorganism...... 122 4.4.1.1 Genomic DNA Isolation, Polymerase Chain Reaction and Transformation of the 16S rRNA Gene ...... 122 4.4.1.2 Analyses of the 16S rRNA Gene ...... 122 4.4.2 Experimental Layout for the Cultivation of Eubacterium limosum and the Subsurface Biome ...... 124 4.4.3 Cultivation and Assessments of Eubacterium limosum ...... 125 4.4.4 Cultivation and Assessments of the Subsurface Biome ...... 129 4.4.5 Microbial Diversity Assessments of the Subsurface Biome using Denaturing Gradient Gel Electrophoresis ...... 137 4.4.5.1 Genomic DNA Isolation and Polymerase Chain Reaction ...... 137 4.4.5.2 Nested Polymerase Chain Reaction ...... 139 4.4.5.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses ...... 140 4.5 CONCLUSIONS ...... 143 4.6 REFERENCES...... 144
Contents iv
CHAPTER 5: DESIGN AND CONSTRUCTION OF THE SYRINGE INCUBATORS AND THE BIOREACTOR ...... 151 5.1 INTRODUCTION ...... 151 5.2 AIMS OF THIS CHAPTER ...... 153 5.3 MATERIALS AND METHODS ...... 154 5.3.1 The Syringe Incubators ...... 154 5.3.1.1 Design and Construction ...... 156 5.3.1.2 Syringe Preparation ...... 159 5.3.1.3 Operational Procedure ...... 159 5.3.2 The Bioreactor ...... 161 5.3.2.1 Designs and Constructions ...... 161 5.3.2.2 Operational Procedures ...... 176 5.4 CONCLUSIONS ...... 178 5.5 REFERENCES...... 179
CHAPTER 6: HIGH PRESSURE STUDIES USING THE SYRINGE INCUBATORS ...... 188 6.1 INTRODUCTION ...... 188 6.1.1 Supercritical Carbon Dioxide ...... 189 6.1.2 Microbial Interaction with Supercritical Carbon Dioxide ...... 190 6.2 AIMS OF THIS CHAPTER ...... 191 6.3 MATERIALS AND METHODS ...... 192 6.3.1 Cultivation Conditions and Analyses for Eubacterium limosum ...... 192 6.3.2 Cultivation Conditions and Analyses for the Subsurface Biome ...... 194 6.3.3 Experimental Layout for the Cultivation of Eubacterium limosum and the Subsurface Biome ...... 195 6.4 RESULTS AND DISCUSSIONS ...... 196 6.4.1 Cultivation and Analyses of Eubacterium limosum ...... 196 6.4.2 Cultivation and Analyses of the Subsurface Biome ...... 201 6.5 CONCLUSIONS ...... 208 6.6 REFERENCES...... 210
CHAPTER 7: CARBON SEQUESTRATION SIMULATIONS OF THE SUBSURFACE BIOME USING A CONTINUOUS HIGH PRESSURE BIOREACTOR ...... 216 7.1 INTRODUCTION ...... 216 7.2 AIMS OF THIS CHAPTER ...... 217
Contents v
7.3 MATERIALS AND METHODS ...... 218 7.3.1 Adhesion Studies ...... 218 7.3.2 Carbon Sequestration Simulations with the Subsurface Biome ...... 219 7.3.3 Bioreactor Control Experiments ...... 221 7.3.4 Whole Transcriptome Sequencing using Ion Torrent ProtonTM ...... 222 7.3.4.1 Total RNA Isolation ...... 222 7.3.4.2 RNA Sequencing and Analyses ...... 222 7.4 RESULTS AND DISCUSSIONS ...... 224 7.4.1 Carbon Sequestration Simulations with the Subsurface Biome ...... 224 7.4.2 Geochemical Analyses ...... 226 7.4.3 X-Ray Fluorescence Analyses ...... 229 7.4.4 Bioreactor Control Experiments ...... 231 7.4.5 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis ...... 232 7.4.5.1 Genomic DNA Isolation and Polymerase Chain Reaction ...... 232 7.4.5.2 Nested Polymerase Chain Reaction ...... 233 7.4.5.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses ...... 234 7.4.6 Microbial Diversity Assessments using Targeted rRNA Gene Sequencing ...... 235 7.4.7 Microbial Diversity Assessments using Visual Methods ...... 238 7.4.7.1 Scanning Electron Microscopy ...... 238 7.4.8 Whole Transcriptome Sequencing using Ion Torrent ProtonTM ...... 241 7.5 CONCLUSIONS ...... 249 7.6 REFERENCES...... 251
CHAPTER 8: FINAL CONCLUSIONS ...... 261
SUMMARY ...... 265
OPSOMMING ...... 267
Contents vi
ACKNOWLEDGEMENTS
This was a very big research project and so many people were involved for whom I wish to express my sincere gratitude and appreciation, in no specific order, because without them, I would not have been able to complete this research.
My promoter, my supervisor and my mentor, Prof. Esta van Heerden, for the immense role she played in making me the scientist I am today, since I started in her laboratory as an undergraduate student twelve years ago, for every opportunity she gave me to help build my career through local and international interactions, for all her guidance and especially her faith in me throughout this study and my career thus far and for her invaluable assistance in finalizing this thesis. I could not have asked for a better mentor and will always be grateful for everything I have learned from her.
My co-promoters, Prof. Derek Litthauer, Prof. Tullis Onstott, Prof. Tommy Phelps and Dr. Tony Surridge, for their valuable support throughout this research, but also for sharing their knowledge with me and for always being willing to help and give advice when needed, not only for this project but every time I turned to them with a question or a problem. I have learned so much from each one of them.
The teams from the Star Diamonds Mine, Petra Diamonds and Frontier Mining, for making a study site available for this research and for allowing me access for the duration of this project, but especially Ben Visser, for accommodating me every time I went to the mine. They were always welcoming and willing to help in any way they could.
My family - my parents, Rassie and Ria Erasmus and my sister and brother-in-law, Elilyia and Morné Shardelow, for all the sacrifices they made so that I can have the opportunity to study, for all their long hours that went into finalizing this script. I will always be grateful for their unfailing love, their endless understanding and support
Acknowledgements vii
and for financial assistance, not only for the time of this study, but throughout my life. My niece and godchild, Hayley Shardelow, and all my four-legged “children” just for being a light in my life.
Everyone in the Departments of Instrumentation and Electronics, who contributed to building and designing the high-pressure equipment used in this research, especially Piet Botes, whom I feel did a magnificent job and always thought about my safety first.
Wally van der Hoven and Sarel Marais, for their tremendous amount of hours and effort that went into the final designs and the construction of the high-pressure bioreactor. So much time, money, struggles and frustrations went into this equipment and they were always willing to help again, each time when we encountered yet another problem. They were always so enthusiastic about solving the problem, which encouraged me to try again until we finally overcame each obstacle.
Dr. Fanie Otto from SASOL, for his contribution towards components used in the bioreactor. High-pressure equipment is very expensive and he saved us a lot of money.
Annalize Visser and WG van der Hoven for all their effort into drawing the official designs for my high-pressure equipment.
Elmarie Prinsloo, for her endless support and encouragement towards Esta and myself, especially during the final days of writing this thesis.
Hanlie Grobler and Prof. Pieter van Wyk from the Centre for Microscopy for all the hours they spent with me in front of the SEM, helping me to obtain some very magnificent images.
Prof. Rob Bragg, for taking the photos of all the high-pressure equipment.
All collaborators, national and international, who helped with specific analyses, but also gave insights into various aspects that facilitated this research project.
Acknowledgements viii
Everyone in the Department of Microbiology, Biochemistry and Food Biotechnology, who shared ideas, made suggestions, helped where they could and for their guidance and interest shown.
All the past and previous members of the Extreme Biochemistry research group who have been part of my family in the lab, as well as all my other friends and family for their friendship, for supporting me through good and bad times, for sharing in my success and failures and for their care and understanding, inside and outside of this department, but especially Jou-An Chen, Elizabeth Ojo, Errol Cason, Marcele Vermeulen, Kay Kuloyo and Arista van der Westhuizen for they were always willing to help me with either sampling and preparation, administrative arrangements, data analyses or any experiment where I needed assistance.
The South African Centre for Carbon Capture and Storage (SACCCS), the National Research Foundation (NRF), the Ernst and Ethel Eriksen Trust, the Technology Innovation Agency (TIA), the TATA group, the Dean from the Faculty of Natural and Agricultural Sciences, the Office for International Affairs at the UFS, the Deep Carbon Observatory (DCO) and Shimadzu for any means of financial support during this study, whether it was for equipment, for travel, for research or for personal funds.
Above all to God, my Creator, my Friend and my Saviour for every opportunity and giving me the ability to do this work and for all His love, grace and guidance during these years and throughout my life. In Deo Confidimus.
Acknowledgements ix
LIST OF SYMBOLS AND ABBREVIATIONS
% Percentage ~ More or less < Less than > Greater than ≥ Greater than/Equal to °C Degrees Celsius °F Degrees Fahrenheit µg.ml-1 Microgram per millilitre µl Microliter µm Micrometre µM Micromolar
µmax Maximum specific growth rate ‰ Per mille/Parts per thousand Acetyl-CoA Acetyl coenzyme A Aer Aerotaxis receptor AF Acetate fermentation AMS Accelerator mass spectrometry APSR Adenylylsulphate reductase Arc Archaea atm Atmosphere ATP Adenosine triphosphate Bac Bacteria BID Barrier discharge ionization BLAST Basic local alignment search tool bp Base pairs BP Before present CA Carbonic anhydrase CAD Computer-aided design
List of Symbols and Abbreviations x
CAE Computer-aided engineering CAF Central analytical facilities CBM Coal bed methane CCS Carbon capture and storage cDNA Complementary deoxyribonucleic acid cm Centimetre cm3 Cubic centimetre CO2CRC Carbon dioxide cooperative research centre CoB-SH Coenzyme B CoM-SH Coenzyme M
CR CO2 reduction CSIR Council for scientific and industrial research CTAB Cetyl trimethylammonium bromide DCO Deep carbon observatory DGGE Denaturing gradient gel electrophoresis dH2O Distilled water DIC Dissolved inorganic carbon DNA Deoxyribonucleic acid dNTPs Deoxyribonucleotide triphosphates DO Dissolved oxygen DOC Dissolved organic carbon DSMZ Deutsche sammlung von mikroorganismen und zellkulturen dsrA Dissimilatory sulphite reductase alpha subunit dsrC Dissimilatory sulphite reductase gamma subunit EC Electrical conductivity ECBM Enhanced coal bed methane ed. Edition EDS Energy dispersive x-ray EDTA Ethylenediaminetetraacetic acid EGR Enhanced gas recovery emPCR Emulsion polymerase chain reaction EOR Enhanced oil recovery EPA United States environmental protection agency EPS Extracellular polymeric substances et al. et alii/and others
List of Symbols and Abbreviations xi
EtBr Ethidium bromide EtOH Ethanol Euk Eukarya FE-SEM Field emission scanning electron microscopy FFKM Perfluoro-elastomers FW Fissure water g Gram g.L-1 Gram per litre G3P Glyceraldehyde-3-phosphate GC Gas chromatography gDNA Genomic deoxyribonucleic acid GFL® Gesellschaft für labortechnik GMWL Global meteoric water line h-1 Per hour HDPE High-density polyethylene HDR Heterodisulphide reductase HPLC High performance liquid chromatograph IC Ion chromatography ICP Inductively coupled plasma ICP-AES Inductively coupled plasma atomic emission spectroscopy IGS Institute for ground water studies in silco Performed on computer/Via computer simulation in situ On-site IPCC Intergovernmental panel on climate change IPTG Isopropyl β-D-1-thiogalactopyranoside IR Infrared ISBN International standard book number Ka Thousand years ago/from the present km Kilometre kPa Kilopascal L Litre L.min-1 Litre per minute LB Luria-bertani LExEn Life in extreme environments LOI Loss on ignition
List of Symbols and Abbreviations xii
LTP Life tree project M Molar m Metre Ma Million years ago/from the present masl Metres above sea level mbs Metres below surface MCP Methyl-accepting chemotaxis protein mg Milligram mg.L-1 Milligram per litre mg.mL-1 Milligram per millilitre MG-RAST Metagenomics RAST ml Millilitre ml.L-1 Millilitre per litre ml.min-1 Millilitre per minute mm Millimetre mM Millimolar MM Minimal medium mm² Square millimetre MPa Megapascal mS.cm-1 Millisiemens per centimetre mS.m-1 Millisiemens per meter mV Millivolts MΩ.cm-1 Mega-ohm per centimetre N Normal/normality N.m-1 Newton metre NADPH Nicotinamide adenine dinucleotide phosphate NASA National aeronautics and space administration NCBI National centre for biotechnology information NDIR Non-dispersive infrared sensor ng Nanogram ng.µl-1 Nanogram per microlitre nm Nanometre NMR Nuclear magnetic resonance NMWC Nominal molecular weight cut-off Myr−1 Per million years
List of Symbols and Abbreviations xiii
NSF National science foundation NZ New Zealand OD Optical density ORP Oxidation reduction potential Pc Critical pressure PCOR Plains carbon dioxide reduction partnership PCR Polymerase chain reaction PES Polyethersulone PGMTM Personal genome machineTM pH Measure of the acidity or basicity of a solution pHi Intracellular pH Pmol Picomole pmol.µl-1 Picomole per microlitre ppm Parts per million PS Polysulfone Pt Platinum PTFE Polytetrafluoroethylene PTP Pico titer plate PVC Polyvinyl chloride PVDF Polyvinylidene fluoride rDNase Recombinant DNase Rh Rhodium RID Refractive index detector RNA Ribonucleic acid rpm Revolutions per minute rRNA Ribosomal ribonucleic acid RSA Republic of South Africa RT Reverse transcription RuBisCO Ribulose-1,5-biphosphate carboxylase/oxygenase SACCCS South African centre for carbon capture and storage SANEDI South African national energy development institute SANERI South African national energy research institute sc-CO2 Supercritical carbon dioxide SD Star Diamonds SEM Scanning electron microscopy
List of Symbols and Abbreviations xiv
SOC Super optimal broth with catabolite repression (SOB with added glucose) SRB Sulphate-reducing bacteria STP Standard temperature and pressure TAE Tris-acetate-ethylenediaminetetraacetic acid Taq DNA polymerase Thermostable deoxyribonucleic acid polymerase named after Thermus aquaticus Tc Critical temperature TCA Tricarboxylic acid TDS Total dissolved solids TE Tris-ethylenediaminetetraacetic acid TEEIC Tribal energy and environmental information clearinghouse TfbI Transformation buffer I TfbII Transformation buffer II TFF Tangential flow filtration Tm Melting temperature of primers TN Total nitrogen TOC Total organic carbon UFS University of the Free State UHP Ultra-high purity UPS Uninterruptible power supply USA United States of America UV Ultraviolet UV/Vis Ultraviolet-visible spectrophotometry V Volts V. Version v/v Volume per volume v/v/v Volume per volume per volume w/v Weight per volume WTW Wissenschaftlich-technische werkstätten x g Acceleration due to gravity X-Gal 5-Bromo-4-chloro-3-indolyl-beta-D- galactopyranosidehosphate XRF X-ray fluorescence analyses
List of Symbols and Abbreviations xv
LIST OF FIGURES
Figure 1.1: Global greenhouse gas emissions by source, based on global emissions from 2004 [Taken from EPA, (2015)]...... 2
Figure 1.2: Natural and human enhanced greenhouse effects [Taken from livescience.com, (2015)]...... 2
Figure 1.3: Global greenhouse gas emissions by gas, based on global emissions from 2004 [Taken from EPA, (2015)]...... 3
Figure 1.4: Direct measurements for the increase in the atmospheric CO2 levels during recent years [Taken from NASA, (2015)]...... 4
Figure 1.5: The change in global surface temperature relative to 1951-1980 average temperatures [Taken from NASA, (2015)]...... 4
Figure 1.6: Potential storage options for CO2 [Adapted from Sims et al., (2007)]...... 6
Figure 1.7: Possible geological storage options for CO2 [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)]...... 7
Figure 1.8: Properties for suitable storage rocks [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)]...... 8
Figure 1.9: Possible ocean storage options for CO2 [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)]...... 8
Figure 1.10: The process of terrestrial sequestration [Taken from tececo.com, (2015)]. . 9
List of Figures xvi
Figure 1.11: The process for mineral carbonation [Taken from decarboni.se, (2015)]. .. 10
Figure 1.12: Global CO2 storage potential, with estimate storage capacities of 2 – 10
trillion tons of CO2 [Taken from stanford.edu, (2015)]...... 11
Figure 1.13: Possible CO2 storage sites for South Africa [Taken from Cloete, (2010)]. . 11
Figure 3.1: Sandstone rock with a clastic texture [Taken from Geology4today, (2015)]. 37
Figure 3.2: The Star Diamonds mine with shaft 4 that was used as the study site for this research...... 40
Figure 3.3: An approximate position of the borehole, intersecting the Karoo sandstone fissure of the Star Diamonds mine (Supplied by Petra Diamonds/Frontier Mining)...... 41
Figure 3.4: The encased borehole, intersecting the Karoo sandstone fissure water, which was used for sampling in the Star Diamonds mine...... 41
Figure 3.5: The organic-free manifold used for sampling, connected to the casing inside the borehole...... 42
Figure 3.6: On-site equipment used for (A) handheld probe measurements, (B) CHEMets® kits, (C) dissolved oxygen, (D) water flow rate, and (E and F) gas flow rate...... 44
Figure 3.7: Methods used for (A) the sterile 0.2 µM IsoporeTM membrane filter that was inserted into a sterile 25 mm in-line Polyacetal Gelman holder, (B) flushing serum vials with nitrogen gas, (C) filtering water through a sterile 0.2 µM syringe filter, and (D) on-site sampling of serum vials...... 47
Figure 3.8: Methods used for (A) evacuating serum vials, (B) collecting gas samples on- site, (C) collecting water samples for gas when the gas flow was too slow, (D) flushing of the copper tubes with nitrogen gas (99.999%), (E) preventing
List of Figures xvii
air from flowing back into the copper tubes, and (F and G) crimp sealing of the copper tubes...... 49
Figure 3.9: The Karoo sandstone obtained from the Star Diamonds mine...... 50
Figure 3.10: Separation of the crushed rock using a 2.8 mm and a 4.7 mm strainer. .... 50
Figure 3.11: Methods used for (A) sterilizing the canister with the outlet valve opened, (B) flushing the canister, (C and D) collecting fissure water on-site, and (E- G) filtering water from the canister...... 53
Figure 3.12: (A) The SterivexTM filters used to (B) collect biofilm on-site through direct connection to a quick connect clip of the sampling manifold...... 54
Figure 3.13: Tangential flow filtration of the fissure water (A) inside the laboratory and (B) inside the mine...... 55
Figure 3.14: (A) A massive filter used for collecting biofilm from fissure water in larger quantities through (B) direct connection to a quick connect clip of the sampling manifold to the inlet side of the filter and (C) the flow accumulator connected to the outlet side of the filter...... 56
Figure 3.15: (A) The cap of the Carboy bottle with one inlet and two outlets with (B) one of the outlets connected to the pump, and (C) the other outlet left open as an overflow, (D) the tube connected on the inside of the cap, and (E) the pump connected to the inlet of the massive filter...... 57
Figure 3.16: (A) The custom made PVC cartridges with its fitting and caps and (B) the cartridges directly connected to the sampling manifold on the borehole. ... 58
Figure 3.17: A Durov diagram for the geochemical analyses of the fissure water...... 75
13 2 Figure 3.18: (A) δ CCH4 and δ HCH4 values for all samples compared to empirically
determined fields for thermogenic gas and microbial gas produced by CO2 reduction (CR) and acetate fermentation (AF) as described in Whiticar,
List of Figures xviii
2 18 (1999) and (B) δ HH2O and δ OH2O for fissure water samples. The global meteoric waterline (GMWL) is from Craig, (1961). In both graphs, samples from the Karoo formation (Middelbult mine) are indicated by (+). All samples from the Witwatersrand Supergroup are open symbols, and all samples from the Ventersdorp Supergroup are solid symbols. Symbols from individual mines are as follows: Merriespruit and Masimong (circles), Beatrix (squares), Evander (triangles), Kloof (diamonds). Kidd Creek data (X) are for abiogenic methane (Sherwood Lollar et al., 2002). (Obtained from Ward et al., 2004). Samples from the Star Diamonds mine are indicated by a red star...... 77
Figure 3.19: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific), used to determine the size of the amplicons, with the amplified rRNA gene fragments for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The samples are shown in lanes 4 – 12...... 79
Figure 3.20: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific), used to determine the size of the amplicons, with the amplified V3/V4 hypervariable regions for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The samples are shown in lanes 4 – 12...... 80
Figure 3.21: DGGE diversity profiles for (A) Archaea, (B) Bacteria, and (C) Eukarya. ... 81
Figure 3.22: MG-RAST taxonomic hit distribution of the diversity in the subsurface biome for (A) domain, (B) phylum, (C) class, and (D) order, with (E) a rarefaction curve indicating the annotated species richness...... 84
Figure 3.23: MG-RAST Krona diagram for the Bacteria at order level...... 85
Figure 3.24: MG-RAST Krona diagram for the Eukarya at order level...... 86
List of Figures xix
Figure 3.25: MG-RAST Krona diagram for the Archaea at order level...... 87
Figure 3.26: Negative stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm...... 90
Figure 3.27: Live/dead stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm...... 91
Figure 3.28: Gram stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm...... 92
Figure 3.29: Scanning electron microscopy images of the fissure water and biofilm from the Star Diamonds mine. Scale bars are equal to either 10 µm or 1 µm. .. 93
Figure 4.1: The pGEM®-T Easy Vector map and sequence reference points (Promega)...... 113
Figure 4.2: (A) A Hungate tube pressurized to 2 bar and (B-D) the syringe-tipped, high pressure hoses used to pressurize the tubes...... 118
Figure 4.3: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, and (B) the amplified 16S rRNA gene fragment from E. limosum. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown and in lanes 2 – 5, the 16S rRNA gene fragment...... 122
Figure 4.4: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the inserts, and (B) the double digestion profile for E. limosum. In lane 1, the O’GeneRulerTM DNA Ladder Mix are shown and in lanes 2 – 5, the ~3000 bp plasmid backbone and the ~1500 bp insert...... 123
Figure 4.5: A summary of the experimental layout for the cultivation of (A) E. limosum and (B) the subsurface biome...... 124
List of Figures xx
Figure 4.6: Growth curves for E. limosum under anaerobic conditions (A) in blue and low pressure (B) in purple (FW = fissure water, +SS = with sandstone, -SS = without sandstone)...... 125
Figure 4.7: Negative stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm...... 127
Figure 4.8: Live/dead stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm...... 128
Figure 4.9: Gram stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm...... 129
Figure 4.10: Growth curves of the subsurface biome under aerobic conditions (C) in orange/yellow (FW = fissure water, +SS = with sandstone, -SS = without sandstone)...... 130
Figure 4.11: Growth curves of the subsurface biome under anaerobic conditions (D) in orange/yellow and low pressure (E) in red/pink (FW = fissure water, +SS = with sandstone, -SS = without sandstone)...... 130
Figure 4.12: Negative stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm...... 134
Figure 4.13: Live/dead stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm...... 135
Figure 4.14: Gram stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm...... 137
Figure 4.15: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, with the amplified rRNA gene fragments
List of Figures xxi
for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. Lanes 4 – 6 contain the LB, FW + SS, and FW - SS from the aerobic growth studies. Lanes 7 – 9 contain the LB, FW + SS, and FW - SS from the anaerobic growth studies. Lanes 10 – 12 contain the LB, FW + SS, and FW - SS from the low pressure growth studies...... 138
Figure 4.16: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons with the amplified V3/V4 hypervariable regions for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The DGGE PCR amplicons are shown in the rest of the lanes...... 139
Figure 4.17: DGGE diversity profiles for (A) Archaea, (B) Bacteria, and (C) Eukarya. Lanes 1 – 3, 8 – 10, and 11 – 13 contain the low pressure growth studies, each time in the order of LB, FW + SS, and FW – SS. Lanes 4 – 6 contain the aerobic growth studies in the order of LB, FW + SS, and FW – SS. Lane 7 contain the anaerobic growth study for LB...... 140
Figure 5.1: Schematic illustration of the microbiological cultivation and incubation technique (Taken from Takai et al., 2008)...... 154
Figure 5.2: Procedure for cultivation in a gas-rich fluid under high hydrostatic pressures (Taken from Takai et al., 2008)...... 155
Figure 5.3: Schematic illustration of the syringes inside the pressure vessel (Taken from Hutton et al., 2001)...... 155
Figure 5.4: Schematic illustrations of (A) the complete syringe incubator, (B) a cut view of the syringe incubator, (C) the lid, (D) the safety pin, and (E) the syringe ring...... 157
List of Figures xxii
Figure 5.5: Images of (A) all the constructed syringe incubators, (B) the individual components of a syringe incubator, (C) the lid, (D) the 5 ml syringe ring, (E) the 25 ml syringe ring, and (F) the safety pin...... 158
Figure 5.6: (A) Pressurizing the syringe incubators using a hydraulic pump and (B) the modified cylinder connected to the hydraulic pump. The white tubing is used for filling the water cylinder...... 160
Figure 5.7: Schematic illustration of the first design of the bioreactor...... 162
Figure 5.8: Schematic illustration of the second design of the bioreactor...... 163
Figure 5.9: Schematic illustration of the third design of the bioreactor...... 164
Figure 5.10: (A) The constructed bioreactor, (B) the bioreactor inside the safety cabinet, and (C) the HPLC pump connected to the bioreactor, with the line to the
CO2 gas cylinder, visible outside of the safety cabinet...... 165
Figure 5.11: Schematic illustration of the fourth design of the bioreactor...... 168
Figure 5.12: The bioreactor, constructed from the fourth design...... 169
Figure 5.13: Schematic illustration of the fifth design of the bioreactor...... 172
Figure 5.14: Schematic illustration of the sixth design of the bioreactor...... 174
Figure 5.15: (A) The bioreactor, constructed from the fifth and sixth designs, (B) a gas tight syringe connected to the sampling port, (C) the reactor, (D) the three- way valves regulating the variable media reservoirs, (E) the three-way valves regulating the piston pressure regulators, (F) the connection to the
CO2 gas cylinder and the media reservoir through the peristaltic pump, and (G) the bioreactor as a whole...... 175
List of Figures xxiii
Figure 6.1: A pressure-temperature phase diagram for carbon dioxide (Taken from https://en.wikipedia.org/wiki/Supercritical_carbon_dioxide)...... 189
Figure 6.2: The prepared syringes for (A) 5 ml without gas, (B) 5 ml with gas, (C) 25 ml without gas, and (D) 25 ml with gas. (The LB medium is displayed in the yellow colour and the PYG medium in the red)...... 192
Figure 6.3: A summary of the experimental layout for the cultivation of E. limosum and the subsurface biome under increasing pressure and (A) different gas concentrations, with (B) different media compositions...... 195
Figure 6.4: The growth of E. limosum after 48 hours in PYG medium, under various high
pressures and 20% CO2...... 196
Figure 6.5: The growth of E. limosum after 48 hours in PYG medium, under various gas concentrations at 70 and 80 bar...... 196
Figure 6.6: The growth of E. limosum after 48 hours in various media compositions,
under 100% CO2 at 70 and 80 bar...... 197
Figure 6.7: Live/Dead stain images of E. limosum after 48 hours in PYG medium, under
various high pressures and 20% CO2. All scale bars are equal to 2 µm. ... 199
Figure 6.8: Live/Dead stain images of E. limosum after 48 hours in PYG medium, under various gas concentrations at 70 and 80 bar. All scale bars are equal to 2 µm...... 200
Figure 6.9: Live/Dead stain images of E. limosum after 48 hours in various media
compositions, under 100% CO2 at 70 and 80 bar. All scale bars are equal to 2 µm...... 201
Figure 6.10: The growth of the subsurface biome after 48 hours in LB medium, under
various high pressures and 20% CO2...... 202
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Figure 6.11: The growth of the subsurface biome after 48 hours in LB medium, under various gas concentrations at 70 and 80 bar...... 202
Figure 6.12: The growth of the subsurface biome after 48 hours in various media
compositions, under 100% CO2 at 70 and 80 bar...... 203
Figure 6.13: Live/Dead stain images of the subsurface biome after 48 hours in LB
medium, under various high pressures and 20% CO2. All scale bars are equal to 2 µm...... 205
Figure 6.14: Live/Dead stain images of the subsurface biome after 48 hours in LB medium, under various gas concentrations at 70 and 80 bar. All scale bars are equal to 2 µm...... 206
Figure 6.15: Live/Dead stain images of the subsurface biome after 48 hours in various
media compositions, under 100% CO2 at 70 and 80 bar. All scale bars are equal to 2 µm...... 207
Figure 7.1: The prepared syringes, used for the adhesion studies...... 218
Figure 7.2: GC analyses of (A) fissure water, before the addition of CO2, (B) after the
addition of CO2, and (C) S8 - after 6 weeks...... 226
Figure 7.3: A Durov diagram of the geochemical analyses of the fissure water and the bioreactor water after six weeks...... 228
Figure 7.4: Comparisons of the geochemical analyses for the bioreactor with the fissure water. The bioreactor displayed an increase in the aluminium, the iron and the M-alkalinity...... 228
Figure 7.5: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, with the amplified rRNA gene fragments for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative
List of Figures xxv
controls, respectively. Lanes 4 – 6 contain the amplicons from the bioreactor...... 233
Figure 7.6: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons with the amplified V3/V4 hypervariable regions for (B) Archaea and (C) Bacteria. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. Lanes 4 – 6 contain the amplicons from the bioreactor...... 234
Figure 7.7: DGGE diversity profiles of (A) Archaea and (B) Bacteria...... 234
Figure 7.8: MG-RAST taxonomic hits distribution of the diversity in the subsurface biome for (A) class and (B) order, with (C) a rarefaction curve indicating the annotated species richness...... 236
Figure 7.9: MG-RAST Krona diagram of the Bacteria at order level...... 237
Figure 7.10: Scanning electron microscopy images of the biome after being subjected to CCS conditions. Scale bars are equal 1 µm...... 238
Figure 7.11: Scanning electron microscopy images, indicating the tube-like and hyphae- like structures. Scale bars are equal to either 1 µm or 10 µm...... 239
Figure 7.12: EDS analyses, confirming the presence of aluminium precipitation...... 240
Figure 7.13: The C1 carbon fixation pathways cycles [Taken from Bar-Even et al., (2012)]...... 242
Figure 7.14: The acetyl-CoA-succinyl-CoA carbon fixation cycles [Taken from Bar-Even et al., (2012)]...... 243
Figure 7.15: MG-RAST Krona diagram of the functional category hits distribution of the subsurface biome...... 246
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Figure 7.16: MG-RAST Krona diagram of CO2 fixation in the subsurface biome...... 247
Figure 7.17: MG-RAST KeggMapper. The genes present within the subsurface biome are shown in the blue highlighted lines...... 248
List of Figures xxvii
LIST OF TABLES
Table 3.1: The universal oligonucleotide primers used for amplification of the Archaeal 16S; Bacterial 16S and Eukaryal 18S rRNA gene fragments...... 60
Table 3.2: The DGGE oligonucleotide primers used for amplification of the V3/V4 hypervariable regions for Archaea; Bacteria and Eukarya respectively. Underlined sequences indicate the GC-clamp...... 61
Table 3.3: The oligonucleotide primers used for the re-amplification of the V3/V4 hypervariable regions for Archaea; Bacteria and Eukarya respectively...... 63
Table 3.4: A summary of all the samplings done at the Star Diamonds mine...... 71
Table 3.5: The collaborators and institutions responsible for the analyses of selected samples...... 72
Table 3.6: All the basic, on-site analyses of the fissure water that was measured over time...... 73
Table 3.7: All the geochemical analyses on the fissure water...... 74
Table 3.8: Major-element XRF analyses of the sandstone, before and after sterilization...... 77
Table 4.1: The RNA polymerase promoter primers used for sequencing...... 115
Table 4.2: BLAST results for the 16S rRNA gene sequence of E. limosum...... 123
Table 7.1: Conditions for the bioreactor control experiments...... 222
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Table 7.2: The pH and growth measurements obtained for the bioreactor during the six weeks...... 224
Table 7.3: Geochemical analyses of the bioreactor...... 227
Table 7.4: Major-element XRF analyses of the sandstone before and after six weeks inside the bioreactor...... 229
Table 7.5: Geochemical analyses for the fissure water and the control experiments. 231
Table 7.6: XRF analyses for the sandstone and the control experiments...... 232
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CHAPTER 1
LITERATURE REVIEW
1.1 GLOBAL CLIMATE CHANGE AND THE GREENHOUSE EFFECT
The average global temperatures are increasing and have resulted in changes in the weather and climate. Global warming is used to describe the current and on-going increase in the average global temperature of the atmosphere near the surface of the Earth and is caused mostly by increasing concentrations of greenhouse gases, in particular carbon dioxide (CO2) in the atmosphere (Vitousek, 1994; Pacala & Socolow, 2004; Ramanan et al., 2009; Solomon et al., 2009; Velea et al., 2009; EPA, 2015; IPCC, 2015; NASA, 2015; SACCCS, 2015; TEEIC, 2015). Global warming represents only one aspect of climate change, since climate change refers to any significant change in measures of climate, for instance temperature, precipitation, or wind patterns which continues for an extended period of time such as decades and even longer (Vitousek, 1994; EPA, 2015). Climate change is occurring from natural factors such as respiration and volcanic eruptions, but ultimately, human activities are the biggest contributor to the recent climate changes.
According to the last available global greenhouse gas emission by source, made by the Intergovernmental Panel on Climate Change (IPCC) in 2007, human activities such as deforestation, transportation, industrial processes, and selected agricultural practices have resulted in the release of large amounts of additional CO2 and other greenhouse gases into the atmosphere, with the leading contributor for these emissions being from the burning of fossil fuels such as coal, natural gas, and oil for power generation and energy supply (Figure 1.1) (Vitousek, 1994; Solomon et al., 2009; Velea et al., 2009; EPA, 2015; SACCCS, 2015; TEEIC, 2015). These additional emissions are responsible for the greenhouse effect (Figure 1.2). Radiation, primarily from the sun, reaches the surface of the Earth in the form of visible light, ultraviolet (UV), infrared (IR) and other forms of radiation. Around 30% of this radiation that reaches the Earth's atmosphere is immediately reflected back out to space through clouds, ice, snow, sand and other reflective surfaces
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(NASA, 2015). The remaining 70% is absorbed by the oceans, land, and atmosphere, thereby, warming up the Earth and heat is released in the form of IR thermal radiation, which is then radiated back into the atmosphere on its way to space (EPA, 2015).
Figure 1.1: Global greenhouse gas emissions by source, based on global emissions from 2004 [Taken from EPA, (2015)].
As the layer of greenhouse gases around the planet grows thicker, more heat is trapped in the atmosphere, less radiates away and the Earth slowly heats up, causing the temperature on the Earth to rise and the climate to change and ultimately results in harmful consequences to human health, the economy, and the ecosystems (Vitousek, 1994; Velea et al., 2009; EPA, 2015; IPCC, 2015; TEEIC, 2015).
Figure 1.2: Natural and human enhanced greenhouse effects [Taken from livescience.com, (2015)]. Chapter 1 2
According to the last available global greenhouse gas emission by gas, made by the IPCC in 2007, these gases include water vapour, CO2, methane (CH4), nitrous oxide (N2O), and fluorinated gases. Of these emissions, CO2 is the most important anthropogenic greenhouse gas (Figure 1.3) (EPA, 2015).
Figure 1.3: Global greenhouse gas emissions by gas, based on global emissions from 2004 [Taken from EPA, (2015)].
Since the beginning of the Industrial Revolution in the early 1800’s, the CO2 levels in the atmosphere have increased by more than 40%, from ~280 parts per million (ppm) in the year 1850 to ~400 ppm today (Figure 1.4), and are at its highest in 650 000 years (Vitousek, 1994; Pacala & Socolow, 2004; Velea et al., 2009; NASA, 2015; SACCCS,
2015). The global CO2 emissions due to the burning of fossil fuels have increased >16 times during 1900 and 2008 (Boden et al., 2010). This has resulted in massive impacts on what is considered normal conditions on the Earth such as changes in rainfall, resulting in floods, droughts, or intensive heat-waves. The oceans are warming and becoming more acidic, ice caps are melting, and sea levels are rising (EPA, 2015; IPCC, 2015; NASA, 2015).
The average temperature of the Earth has increased by 1.4°F (~0.8°C) over the last 100 years (Figure 1.5), where nine of the ten warmest years occurred since the year 2000, with 2014 ranked as the warmest year on record.
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Figure 1.4: Direct measurements for the increase in the atmospheric CO2 levels during recent years [Taken from NASA, (2015)].
Figure 1.5: The change in global surface temperature relative to 1951-1980 average
temperatures [Taken from NASA, (2015)].
Predictions are that the temperatures will increase with another 2 - 11°F over the next century and that these perceived small changes can result potentially dangerous shifts in the climate and weather (NASA, 2015). Each September, the Arctic sea ice reaches its minimum and is now declining at a rate of 13.3% per decade, relative to the 1981 to 2010 average (NASA, 2015).
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Since 2002, Antarctica has been losing about 134 billion metric tons of ice per year, while the Greenland ice sheet has been losing an estimated 287 billion metric tons per year (NASA, 2015). The rise in sea levels are mainly caused by two global warming related factors which are the added water from melting land ice and the expansion of sea water as it warms and the global average sea level has risen by ~178 mm during the past century (NASA, 2015).
Carbon dioxide can stay in the atmosphere for almost a century, therefore, the Earth will continue to heat up in future decades (Solomon et al., 2009; EPA, 2015). As these and other changes become more prominent, humanity and the environment will be facing more severe challenges. Consequently, active carbon management is essential to control the increase in global CO2 emissions and thereby mitigate the impact of climate change by enacting policies that essentially reduce the concentration of CO2 in the atmosphere (Pacala & Socolow, 2004; Surridge & Cloete, 2009; Morozova et al., 2010; EPA, 2015). A possible solution is to replace the use of fossil fuels such as oil, coal, and gas with energy sources like solar, wind and possibly nuclear and use them more efficiently, but the first steps are to address the existing fossil fuel base infrastructure and the excessive levels of
CO2 being emitted globally. Therefore, sequestering anthropogenic CO2 from the atmosphere may allow some time to make the transition to low carbon-emitting technologies (Metz et al., 2007; Sherwood Lollar & Ballentine, 2009; Surridge & Cloete, 2009; Cloete, 2010).
1.2 CARBON SEQUESTRATION
Carbon sequestration, also known as carbon capture and storage (CCS) is a process where CO2 is stored away from the atmosphere, as a means to mitigate global climate change (Metz et al., 2007; Holloway, 2007; Cunningham et al., 2009; Morozova et al., 2010; Glossner, 2013; Mu et al., 2014) and has the potential to store millions of tonnes of
CO2. In order to mitigate the CO2 emissions on a scale that can stabilize the atmospheric greenhouse gases and make an impact on climate change, storage on this scale is necessary. Consequently, CCS has the potential to help meet emission reduction goals (Metz et al., 2007; Holloway, 2007; Surridge & Cloete, 2009; Peters et al., 2011).
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Several possibilities exist for the potential storage of CO2 (Figure 1.6), and include direct/artificial and indirect/natural sequestration. During direct/artificial sequestration, the
CO2 is captured before being released into the atmosphere, and stored through geological sequestration or ocean sequestration. For indirect/natural sequestration, terrestrial or biological sequestration is used, and involves the removal of the CO2 from the atmosphere through plants, trees and subsequent storage in soil. CO2 can also be fixed into inorganic carbonates in the subsurface, through mineral sequestration/carbonation (Metz et al., 2007; Cloete, 2010; EPA, 2015; PCOR, 2015; TEEIC, 2015).
Terrestrial sequestration
Figure 1.6: Potential storage options for CO2 [Adapted from Sims et al., (2007)].
1.2.1 Geological Sequestration
During geological sequestration (Figure 1.7), anthropogenic CO2 is injected into underground formations such as depleted oil and gas fields, un-minable coal beds, and deep saline formations, either onshore or offshore (Metz et al., 2007; Cloete, 2010; TEEIC,
2015). During storage in depleted oil and gas fields, the CO2 is trapped in the pore spaces of the sedimentary rock, leading to enhanced oil and gas recovery (EOR and EGR). For the un-minable coal beds, the CO2 displaces the coal bed methane (CBM) by adsorption onto the coal surfaces, leading to enhanced CBM recovery (ECBM) (Cloete, 2010).
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Storage of CO2 in deep saline formations is generally expected to take place at depths of >800 metres below surface (mbs), or where the ambient pressure and temperature conditions will result in the CO2 being in a supercritical state, thereby filling the pore spaces of the sedimentary rock (Benson & Cole, 2008; Cunningham et al., 2009; Cloete, 2010; Nondorf et al., 2011; Glossner, 2013; Gulliver, 2014).
Figure 1.7: Possible geological storage options for CO2 [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)].
During storage in deep saline formations, the first step is capturing the CO2, generated from human activity and from large industrial facilities such as power plants, oil refineries or cement plants. The captured CO2 is separated from other gases and pressurized to a supercritical fluid. This fluid is then transported to its destination, either through the use of pipelines, trucks, or ships, and is injected deep below the surface of the Earth into porous rock such as sandstone, which has the necessary porosity and permeability (Figure 1.8). This rock should be overlaid with an impermeable cap rock such as shale, which acts as a seal to prevent CO2 from migrating back to the surface of the Earth and ultimately entering the atmosphere (Metz et al., 2007; Holloway, 2007; Cunningham et al., 2009; Cloete, 2010; Glossner, 2013; Gulliver, 2014; TEEIC, 2015).
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Figure 1.8: Properties for suitable storage rocks [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)].
1.2.2 Ocean Sequestration
During ocean sequestration (Figure 1.9), anthropogenic CO2 can potentially be injected into the water column, at >1 000 mbs, through the use of a fixed pipeline or a moving ship or by depositing it onto the sea floor, at >3 000 mbs, through the use of a fixed pipeline or an offshore platform (Metz et al., 2007).
Figure 1.9: Possible ocean storage options for CO2 [Taken from the Carbon dioxide Cooperative Research Centre, (2015) (CO2CRC.com)]. Chapter 1 8
At >1 000 mbs, the CO2 will be able to dissolve and disperse to form part of the global carbon cycle where it will eventually equilibrate with the CO2 in the atmosphere. At >3 000 mbs, the CO2 is denser than water and should form a lake that would delay the dissolution of the CO2 into the surrounding environments (Metz et al., 2007).
1.2.3 Terrestrial Sequestration
Terrestrial sequestration (Figure 1.10) is the natural process of CO2 sequestration that occurs with trees, plants, animals, the oceans, and soil (PCOR, 2015; TEEIC, 2015).
Figure 1.10: The process of terrestrial sequestration [Taken from tececo.com, (2015)].
Trees and plants can use atmospheric CO2 to live and grow through photosynthesis.
During this process, the CO2 becomes part of the plant and may result in carbon that is sequestered for a long period of time in stems and roots, as well as in the soil. When the plant or tree dies, the plant material decomposes, releasing some of the carbon into the atmosphere and capturing the rest within the soil which increases the soil's organic matter content. Terrestrial sequestration can be increased through forest management, reforestation, and afforestation (PCOR, 2015; TEEIC, 2015).
Chapter 1 9
1.2.4 Mineral Sequestration
Mineral sequestration/carbonation (Figure 1.11) is a chemical process of CO2 storage wherein the atmospheric CO2 reacts with metal oxides such as calcium oxide (CaO) or magnesium oxide (MgO) to produce stable carbonate minerals. These oxides are found in silicate minerals or industrial waste streams. This is naturally a very slow process, but can be enhanced through pre-treatment of the minerals (Metz et al., 2007; Cloete, 2010).
Figure 1.11: The process for mineral carbonation [Taken from decarboni.se, (2015)].
1.3 CARBON SEQUESTRATION IN SOUTH AFRICA
Possible carbon sequestration sites exist worldwide, but these storage formations are not evenly distributed (Figure 1.12), therefore, the potential for underground storage of CO2 varies for each country. During preliminary investigations by the Council for Scientific and Industrial Research (CSIR) in 2004, it was determined that South Africa has various sites
(Figure 1.13) for the storage of CO2 and that of the more than 400 million tonnes of annual carbon dioxide emissions in South Africa ~60% can be sequestered (Surridge & Cloete, 2009; Cloete, 2010).
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Figure 1.12: Global CO2 storage potential, with estimate storage capacities of 2 – 10 trillion
tons of CO2 [Taken from stanford.edu, (2015)].
Figure 1.13: Possible CO2 storage sites for South Africa [Taken from Cloete, (2010)].
During 2007, a long-term mitigation scenario study was conducted to determine all the actions that South Africa could undertake to limit the emissions of greenhouse gases. During 2009, an investigation into the regulation gaps for CCS was initiated by the Department of Energy and in In March 2009, the South African Centre for Carbon Capture
Chapter 1 11
and Storage (SACCCS) was established with the support of South African National Energy Development Institute (SANEDI), previously known as the South African National Energy
Research Institute (SANERI) and in 2010, a detailed Atlas on geological storage of CO2 in South Africa was released (Surridge & Cloete, 2009; Cloete, 2010; SACCCS, 2015). This
Atlas indicated that South Africa’s CO2 emissions will increase until 2020 - 2025, stabilize for a decade and then decrease dramatically after 2030 – 2035 (Cloete, 2010).
1.4 THE CARBON CYCLE
The carbon cycle is the circulation and transformation of carbon back and forth between living material and the environment, which includes the terrestrial biosphere, the oceans, the atmosphere, and the geosphere (Detwiler & Hall, 1987; Houghton et al., 1995; Sedjo, 2001; Graber, 2011; TEEIC, 2015).
During carbon cycling between the terrestrial biosphere and the atmosphere, terrestrial plants remove carbon from the atmosphere through photosynthesis, where the CO2 is stored and used for growth. Animals can absorb this carbon through ingestion of plants, where the carbon from nutrients and food is consumed and CO2 is released as waste back into the atmosphere through respiration. Plants can then again absorb this CO2 or microorganisms can break down the animal manure into methane and CO2. This is similar to microorganisms degrading dead plants and animals and converting the carbon into CO2 and methane that becomes part of the soil. Over long periods of time, this dead organic matter is compacted and will become part of the geosphere through a process called sedimentation, the process responsible for the production of fossil fuels such as oil and coal (Detwiler & Hall, 1987; Houghton et al., 1995; Sedjo, 2001; Graber, 2011; TEEIC, 2015).
Carbon cycles between the atmosphere and the ocean through a process called diffusion and aquatic plant life absorbs carbon through photosynthesis. Marine life can absorb this carbon through consumption, which is then returned to the water in air through respiration. When these organisms die, they decay and release carbon into the ocean which can be transferred to the ocean floor through sedimentation. Carbon enters the oceans from the geosphere in the form of bicarbonate. Bicarbonate also forms in the oceans when the atmospheric CO2 dissolves in the water. Some of the marine life can use this bicarbonate to build their skeletons or it can form limestone. The limestone sinks to the bottom of the
Chapter 1 12
ocean, sequestering the carbon in the geosphere for a long time. As ocean waters flows between the cold poles and the equator, and as it cycles to and from the warm shallow waters and the cold deep water, it absorbs and releases carbon from the atmosphere, typically as CO2 (Detwiler & Hall, 1987; Houghton et al., 1995; Sedjo, 2001; Graber, 2011; TEEIC, 2015).
Several processes are involved in cycling carbon between the geosphere and the atmosphere such as volcanic eruptions and the production of cement. Cement is made from limestone removed from the geosphere, and involves heating this limestone, thus releasing its stored carbon as CO2. The burning of fossil fuels also releases carbon into the atmosphere (Detwiler & Hall, 1987; Houghton et al., 1995; Sedjo, 2001; Graber, 2011; TEEIC, 2015).
1.5 THE DEEP SUBSURFACE AND CARBON CYCLING
The terrestrial subsurface biosphere has been estimated to contain 40–50% of the world’s biomass in the form of microorganisms, extending deeper than four kilometres into the Earth’s crust, with cell concentrations ranging from 1x109 to 28 cells/gram or cells/ml (Zhang et al., 2005; Wang et al., 2007; Chivian et al., 2008; Borgonie et al., 2011; Magnabosco et al., 2014; Mu et al., 2014; Rajala et al., 2015). However, comprehensive studies on the phylogenetic diversity and distribution are geographically limited as a result of the vast capacity of the subsurface ecosystems, but also due to the difficulty in obtaining samples from these extreme and highly reducing environments. Thus, the diversity, abundance and function of these subsurface microorganisms remains largely unknown (Ghiorse & Wilson, 1988; Gold, 1992; Parkes et al., 1994; White et al., 1998; Whitman et al., 1998; Pedersen, 2000; Teske, 2005; Chivian et al., 2008; Fry et al., 2008; Ragon et al., 2013; Rajala et al., 2015).
Even though the terrestrial subsurface includes a substantial fraction of the global biosphere (Pedersen, 1993; Onstott et al., 1998; Whitman et al., 1998), these microorganisms are likely to be inactive or have exceptionally slow metabolisms, as they are living under extreme energy limiting conditions, especially due to the low nutrient availability in subsurface environments (Jørgensen & D’Hondt, 2006; Jørgensen & Boetius, 2007; Sogin et al., 2010; Rajala et al., 2015). Geochemical evidence suggests
Chapter 1 13
that these in situ microbial metabolisms are estimated to be in the nano- to micromolar range per year, with an average turnover time of decades or centuries (Phelps et al., 1994; Onstott et al., 2009). Excluded from the presence of sunlight, the deep subsurface provides a natural environment where energy-yielding processes such as radiolytic decomposition of water to yield H2, O2, and H2O2, with the subsequent oxidation of minerals containing reduced forms of sulphur, iron, manganese, or even arsenic may sustain carbon cycling (Lovley & Chapelle, 1995; Onstott et al., 2006; Sogin et al., 2010). Thus, an increase in the metabolic activities of the subsurface microorganisms may occur through the accessibility of chemical energy at sedimentary and/or geochemical interfaces, and is usually derived from redox reactions involving organic matter or inorganic electron donors such as iron, hydrogen, methane, and hydrogen sulphide (Chapelle et al., 2002; Nealson et al., 2005; Onstott et al., 2006; Jorgensen & Boetius, 2007; Ragon et al., 2013).
It was found that H2 might be the most important energy source for these subsurface communities, since subsurface microbial communities make use of substrates, obtained from geochemical processes and not from photosynthetically derived organic carbon (Stevens & McKinley, 1995; Pedersen, 2001; Lin et al., 2005; Nealson et al., 2005; Onstott et al., 2006). Studies conducted on the biogeochemistry of the deep subsurface indicate that, even though all three domains of life (Archaea, Bacteria and Eukarya) have been found to be present in the subsurface, it is generally dominated by the Bacteria, especially the Proteobacteria and the Firmicutes, and that the leading microbial activities in these extreme environments are methanogenesis, sulphate reduction, and fermentation, the terminal steps in the degradation of organic compounds in the carbon cycle (Banfield et al., 1999; Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013; Lau et al., 2014; Magnabosco et al., 2014). However, the role of the microbial communities in the carbon cycling occurring in the subsurface, remains poorly understood and are only minimally represented in carbon cycle models (Sogin et al., 2010; Rajala et al., 2015).
Therefore, an important aspect to consider during carbon sequestration, is the potential response of the subsurface microbial communities towards elevated CO2 levels, since these microorganisms can play a significant role in the carbon cycling that occurs in the subsurface (Chapelle et al., 2002; Morozova et al., 2010; Bordenave et al., 2012; Mu et al., 2014). Most of the prokaryotic microbial communities contain individuals that can grow
Chapter 1 14
autotrophically, thus, contribute to a supply of organic carbon for heterotrophic microorganisms. Even though the Calvin cycle is known to be the most important autotrophic carbon fixation pathway, it is restricted to organisms with high-energy yield from a chemotrophic or phototrophic existence (Hügler et al., 2005). Therefore, the subsurface microorganisms could utilize different CO2 fixation pathways such as the reductive tricarboxylic acid (TCA) cycle, the reductive acetyl-CoA pathway, and the 3- hydroxypropionate cycle (Hügler et al., 2005).
Heterotrophic microorganisms cannot fix carbon; thus, they use organic carbon for growth (Hogg, 2013) and obtain their energy from the oxidation of organic compounds such as lipids, proteins, and carbohydrates, especially glucose. Biologic oxidation of these organic compounds results in the synthesis of Adenosine triphosphate (ATP). Heterotrophic metabolism includes respiration and fermentation (Jurtshuk, 1996; Hogg, 2013). During respiration, the organic compounds are completely oxidized to CO2 and H2O. For aerobic respiration, molecular O2 is the terminal electron acceptor, whereas with anaerobic
- 2- respiration, NO3 , SO4 , CO2, or fumarate can serve as the terminal electron acceptors, depending on the microorganism. Fermentation requires an organic compound such as glucose as a terminal electron or hydrogen acceptor and the formation of end products result from anaerobic dissimilation of organic compounds. Energy, in the form of ATP, is generated through dehydrogenation reactions through the enzymatic breakdown of glucose (Jurtshuk, 1996; Hogg, 2013).
Autotrophic microorganisms can make use of CO2 as their sole carbon source and include the photoautotrophs, which can use light as their energy source and CO2 as the carbon source, as well as the chemoautotrophs (also known as the chemolitotrophs or the chemolitoautotrophs), which use inorganic chemicals as their energy source and CO2 as the carbon source (Jurtshuk, 1996; Lim, 2003; Hogg, 2013). Through a process called photophosphorylation, the photoautotrophs can convert light energy to chemical energy (ATP) during photosynthesis. The chemoautotrophs can remove electrons from inorganic compounds and through using the electron transport chain, reduced nicotinamide adenine dinucleotide phosphate (NADPH) is formed as the end product (Jurtshuk, 1996; Lim, 2003;
Hogg, 2013). The resulting ATP and NADPH can be used for CO2 fixation with the Calvin cycle.
Chapter 1 15
Bacterial methanogenesis can take place without the presence of sunlight by a group of microorganisms, called the methanogens. These microorganisms are thought to be responsible for contributing towards greenhouse gas emissions as they produce methane, for example, in cow manure, since they are abundant in these anoxic habitats (Koide,
1999). The methanogens use CO2 as the carbon source for growth and as a final electron acceptor to produce methane and fix CO2 using the reductive acetyl-CoA pathway. Thus, if
CO2 is injected into the deep subsurface and dissolves in groundwater, the methanogens can reduce the CO2 to produce methane, when H2 is available (Koide, 1999; Ju et al.,
2008; Glossner, 2013). Methane is less soluble in water than CO2 and can easily migrate to Earth’s surface. Therefore, if no cap rock or seal exists to trap the methane in the subsurface, it will be emitted into the atmosphere and contribute to climate change (Koide, 1999).
Chapter 1 16
1.6 CONCLUSIONS
Climate change is predominantly caused by increasing concentrations of greenhouse gases, more specifically CO2, in the atmosphere. Human activities, mainly from the burning of fossil fuels, are the biggest contributor to the increase in the average global temperature of the Earth's atmosphere and to limit these CO2 emissions, carbon resources have to be managed more effectively.
As a possible solution, storage of anthropogenic CO2 into geological formations, away from the atmosphere, may allow some time for the world to transition to renewable energy.
However, CO2 storage sites require that intensive studies before, during, and after injection should be done since there are several potential risks during carbon sequestration if these activities are not monitored.
South Africa has determined that various potential sites for the storage of CO2 are available and that of the more than 400 million tonnes of annual carbon dioxide emissions in South Africa, ~60% can be sequestered. However, during the past few years, there has been an increased understanding of the volume and activities of life in the deep subsurface. The coupled hydrological, microbiological and geochemical processes occurring within these extreme environments may result in natural chemical fluxes that could influence water quality. The deep terrestrial biosphere in South Africa is dominated by radiogenic noble gases and crustal-derived carbon sources, with microbial metabolisms dominated by methanogenesis, sulphate reduction, and fermentation.
The terrestrial subsurface biosphere remains an extreme environment where microbial communities evolved various mechanisms in order to survive the high pressures and temperatures, the limited energy and nutrient sources, the extreme acidity and alkalinity, the metal toxicity, as well as radioactive processes encountered in the subsurface. Therefore, these microorganisms not only have the ability to adapt to these conditions, but they can also change their surrounding environment in order to provide the necessary nutrients to sustain them. As a result, their survival depends mainly on their ecological and metabolic capabilities (Pikuta et al., 2007; Reith, 2011).
Microbial activity in subsurface environments could potentially play a critical role in the cycling of carbon and other elements between the deep and shallow environments on
Chapter 1 17
Earth (Sogin et al., 2010). Consequently, the metabolic activity of these subsurface microbial communities and their response to varying environmental conditions could contribute to substantial changes in the structure and chemical composition of the rock formations by influencing the porosity and permeability of the storage reservoir. These changes can result in possible leakage of the CO2, or even methane, since the CO2 may be fixed into organic matter by chemoautotrophs and in addition converted into methane by the methanogens.
Chapter 1 18
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CHAPTER 2
INTRODUCTION TO PRESENT STUDY
2.1 ABSTRACT
Climate change is taking place world-wide due to the already high levels of greenhouse gases in the atmosphere, specifically CO2, and this is mainly due to anthropogenic inputs such as the global use of fossil fuels and other human or industrial activities (Vitousek, 1994; Pacala & Socolow, 2004; Ramanan et al., 2009; Solomon et al., 2009; Velea et al.,
2009; EPA, 2015; SACCCS, 2015). In the past, little consideration was given to CO2 as an undesirable emission, but due to an atmospheric increase from 280 ppm in 1850 to 400 ppm currently (NASA, 2015; SACCCS, 2015), it has moved to the top of the list for the most undesirable emission on the international climate change agenda (Surridge & Cloete, 2009). Due to industrialization and the growth in population around the world, the use of fossil fuels is predicted to increase, resulting in a corresponding increase in CO2 emissions to the atmosphere (Solomon et al., 2009; Surridge & Cloete, 2009), and this will result in several long-term, undesirable and unmanageable impacts on the economy, the ecology and human health and therefore, carbon resources have to be managed more effectively. This is especially important for emerging countries with coal based energy such as South Africa where coal is the majority of South Africa’s main energy supply and the demand is projected to keep increasing (Surridge & Cloete, 2009; Cloete, 2010).
As a means to reduce anthropogenic CO2 emissions, South Africa is investigating the possibility of CO2 storage in geological structures while developing alternative renewable and/or nuclear energies that could replace the use of fossil fuel based energy (Sherwood Lollar & Ballentine, 2009; Surridge & Cloete, 2009; Cloete, 2010). During geological sequestration, the captured CO2 will be injected several hundreds of metres below the surface of the Earth to porous rocks such as sandstone, where the CO2 will be in a supercritical state (Cunningham et al., 2009; Cloete, 2010; Nondorf et al., 2011). The molecular phylogenetic surveys performed by members of the Deep Carbon Observatory (DCO) Deep Life research program and the Life in Extreme Environments (LExEn)
Chapter 2 28
research program, funded by the National Science Foundation (NSF), have demonstrated that the South African and other subsurface biomes, exactly where CO2 will be stored, harbour highly diverse and novel microbial communities and the metabolic activities of these subsurface biomes is closely tied to the geochemical and mineralogical processes (Stevens et al., 1993; Phelps et al., 1994; Fredrickson & Onstott, 1996; Colwell et al., 1997; Onstott et al., 1998; Pedersen et al., 2000; Moser et al., 2003; Kieft et al., 2005; Gihring et al., 2006; Onstott et al., 2009; Borgonie et al., 2011; Ragon et al., 2013; Lau et al., 2014; Rajala et al., 2015). As a result, sequestering CO2 could influence these microorganisms on a diversity and metabolic level, but to date, studies on subsurface microbial communities have taken a back seat to geological approaches within this research field.
Consequently, predicting the potential impact of long term CO2 sequestration on the subsurface biome and their contribution to carbon cycling is impossible without an experimental knowledge base. However, the first pilot CO2 storage project experiment for South Africa is planned for 2017, with a demonstration plant planned for 2020, and then the commercial operations are planned for 2025 (SACCCS, 2015). Thus, in order to understand the microbial interaction with the CO2, as well as the rock-microbial interactions that might influence the storage process, bench-scale simulations of these interactions was selected for this research, with the use of specifically designed, high pressure equipment. These are essential questions, not only with respect to carbon sequestration but also to study the effect that the storage will have on current subsurface communities and their role in carbon cycling and ultimately subsurface biogeochemistry.
2.2 POSSIBLE OCCURENCES DURING CARBON SEQUESTRATION
As described by Basava-Reddi, (2012), predictions are that if CO2 is injected into the subsurface where microorganisms are present, it is likely to cause an initial decrease in cell numbers as many microorganisms may not survive the initial conditions. However, once injection is completed, a gradual increase in numbers may occur as some of the microorganisms may adapt to these extreme conditions and utilise the CO2 to produce the necessary organic compounds as part of biotic cycling.
The injected CO2 could also affect physicochemical parameters, like the pH, which in turn, will also affect microbial communities. The extent to which this will occur is dependent on
Chapter 2 29
the biome present, the geology of the storage rock, as well as the chemistry of the water
(Basava-Reddi, 2012). Microorganisms will interact with the sequestered CO2 and their activity can potentially cause direct effects, including mineral formation or degradation, changes in the porosity of the storage rock through biofilm formation, and corrosion or indirect effects, including changes in the pH and redox conditions through cycling (Basava- Reddi, 2012). This might result in either positive or negative effects on the storage formations. Potential positive effects include biofilm formation that can help trap the CO2 in the storage rock, thus increasing storage security, as well as enhanced solubility and mineral trapping. Potential negative effects might include enhancing or selecting for methanogens which can lead to possible enhanced methane generation, changes in the porosity of the rock which will affect the storage capacity, and could result in potential methane leakage to the surface (Morozova et al., 2010; Basava-Reddi, 2012; Mu et al., 2014).
Chapter 2 30
2.3 REFERENCES
Basava-Reddi, L. (2012). Microbial Effects on CO2 Storage. IEAGHG Report.
Borgonie, G., García-Moyano, A., Litthauer, D., Bert, W., Bester, A., van Heerden, E., Möller, C., Erasmus, M. and Onstott, T.C. (2011). Nematoda from the Terrestrial Deep Subsurface of South Africa. Nature. 474:79-82.
Cloete, M. (2010). Atlas on Geological Storage of Carbon Dioxide in South Africa. Council for Geoscience.
Colwell, F.S., Onstott, T.C., Dilwiche, M.E., Chandler, D., Fredrickson, J.K., Yao, Q., McKinley, J.P., Boone, D.R., Griffiths, R., Phelps, T.J., Ringelberg, D., White, D.C., LaFreniere, L., Balkwill, D., Lehman, R.M., Konisky, J. and Long, P.E. (1997). Microorganisms from Deep, High Temperature Sandstones: Constraints on Microbial Colonization. FEMS Microbiol Rev. 20:425-435.
Cunningham, A.B., Gerlach, R., Spangler, L. and Mitchell, A.C. (2009). Microbially
Enhanced Geologic Containment of Sequestered Supercritical CO2. Energy Procedia. 1:3245-3252.
Fredrickson, J.K. and Onstott, T.C. (1996). Microbes Deep Inside the Earth. Sci Am. 275:68-73.
Gihring, T.M., Moser, D.P., Lin, L., Davidson, M., Onstott, T.C., Morgan, L., Milleson, M., Kieft, T.L., Trimarco, E., Balkwill, D.L. and Dollhopf, M.E. (2006). The Distribution of Microbial Taxa in the Subsurface Water of the Kalahari Shield, South Africa. Geomicrobiol. J. 23:415–430.
http://climate.nasa.gov/. (2015).
http://www.epa.gov/climatechange/. (2015).
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Kieft, T.L., McCuddy, S.M., Onstott, T.C., Davidson, M., Lin, L., Mislowack, B., Pratt, L., Boice, E., Sherwood Lollar, B., Lippmann-Pipke, J., Pfiffner, S.M., Phelps, T.J., Gihring, T., Moser, D. and van Heerden, A. (2005). Geochemically Generated, Energy-Rich Substrates and Indigenous Microorganisms in Deep, Ancient Groundwater. Geomicrobiol. J. 22:325–335.
Lau, M.C.Y., Cameron, C., Magnabosco, C., Brown, C.T., Schilkey, F., Grim, S., Hendrickson, S., Pullin, M., Sherwood Lollar, B., van Heerden, E., Kieft, T.L. and Onstott, T.C. (2014). Phylogeny and Phylogeography of Functional Genes Shared Among Seven Terrestrial Subsurface Metagenomes Reveal N-Cycling and Microbial Evolutionary Relationships. Front. Microbiol. 5:531.
Morozova, D., Wandrey, M., Alawi, M., Zimmer, M., Vieth, A., Zettlitzer, M and Würdemann, H. (2010). Monitoring of the Microbial Community Composition in
Saline Aquifers during CO2 Storage by Fluorescence in situ Hybridisation. International Journal of Greenhouse Gas Control. 4:981-989.
Moser, D.P., Onstott, T.C., Fredrickson, J.K., Brockman, F.J., Balkwill, D.L., Drake, G.R., Pfiffner, S.M., White, D.C., Takai, K., Pratt, L.M., Fong, J., Sherwood Lollar, B., Slater, G., Phelps, T.J., Spoelstra, N., DeFlaun, M., Southam, G., Welty, A.T., Baker, B.J. and Hoek, J. (2003). Temporal Shifts in the Geochemistry and Microbial Community Structure of an Ultradeep Mine Borehole Following Isolation. Geomicrobiol. J. 20:517–548.
Mu, A., Boreham, C., Leong, H.X., Haese, R.R. and Moreau, J.W. (2014). Changes
in the Deep Subsurface Microbial Biosphere Resulting from a Field-Scale CO2 Geosequestration Experiment. Frontiers in Microbiology. 5:209.
Nondorf, L., Gutierrez, M. and Plymate, T.G. (2011). Modelling Carbon Sequestration Geochemical Reactions for a Proposed Site in Springfield, Missouri. Environmental Geosciences. 18:91-99.
Onstott, T.C., Colwell, F.S., Kieft, T.L., Murdoch, L. and Phelps, T.J. (2009). New Horizons for Deep Subsurface Microbiology. Microbe Magazine. 4(11):499–505.
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Onstott, T.C., Phelps, T.J., Kieft, T., Colwell, F.S., Balkwill, D.L., Fredrickson, J.K. and Brockman, F.J. (1998). In: Enigmatic Microorganisms and Life in Extreme Environments. Springer Science & Business Media Publishers. ISBN: 9780792354925.
Pacala, S. and Socolow, R. (2004). Stabilization Wedges: Solving the Climate Problem for the Next 50 Years with Current Technologies. Science. 305:968-972.
Pedersen, K., Motemedi, M., Karnland, O. and Sandén, T. (2000). Cultivability of Microorganisms Introduced into a Compacted Bentonite Clay Buffer under High-Level Radioactive Waste Repository Conditions. Eng Geol. 58:149-161.
Phelps, T.J., Murphy, E.M., Pfiffner, S.M. and White, D.C. (1994). Comparison between Geochemical and Biological Estimates of Subsurface Microbial Activities. Microb. Ecol. 28:335-349.
Ragon, M., Van Driessche, A.E.S., García-Ruíz, J.M., Moreira, D. and López- García, P. (2013). Microbial Diversity in the Deep-Subsurface Hydrothermal Aquifer Feeding the Giant Gypsum Crystal-Bearing Naica Mine, Mexico. Frontiers in Microbiology. 4:37.
Rajala, P., Bomberg, M., Kietäväinen, R., Kukkonen, I., Ahonen, L., Nyyssönen, M. and Itävaara, M. (2015). Rapid Reactivation of Deep Subsurface Microbes in the Presence of C-1 Compounds. Microorganisms. 3:17-33.
Ramanan, R., Kannan, K., Sivanesan, S.D., Mudliar, S., Kaur, S., Tripathi, A.K. and Chakrabarti, T. (2009). Bio-sequestration of Carbon Dioxide using Carbonic Anhydrase Enzyme Purified from Citrobacter freundii. World J Microbiol Biotechnol. 25:981-987.
Sherwood Lollar, B. and Ballentine, C.J. (2009). Insights Into Deep Carbon Derived from Noble Gases. Nature Geoscience. 2:543-547.
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Solomon, S., Plattner, G., Knutti, R. and Friedlingstein, P. (2009). Irreversible Climate Change due to Carbon Dioxide Emissions. Proc. Natl. Acad. Sci. 106(6):1704-1709.
Stevens, T.O., McKinley, J.P. and Fredrickson, J.K. (1993). Bacteria Associated with Deep, Alkaline, Anaerobic Groundwaters in Southeast Washington. Microb Ecol. 25:35-50.
Surridge, A.D. and Cloete, M. (2009). Carbon Capture and Storage in South Africa. Energy Procedia. 1:2741-2744.
Velea, S., Dragos, N., Serban, S., Ilie, L., Stalpeanu, D., Nicoara, A. and Stepan, E. (2009). Biological Sequestration of Carbon Dioxide from Thermal Power Plant Emissions, by Absorbtion in Microalgal Culture Media. Romanian Biotechnological Letters. 14(4):4485-4500.
Vitousek, P.M. (1994). Beyond Global Warming: Ecology and Global Change. Ecology. 75(7):1861-1876.
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CHAPTER 3
SAMPLING, CHARACTERIZATION AND MICROBIAL DIVERSITY OF THE SITE DESIGNATED TO MIMIC CARBON SEQUESTRATION CONDITIONS
3.1 INTRODUCTION
The CO2 captured and compressed during the geological CCS process is sequestered several hundreds of metres below the surface of the Earth in porous rocks such as sandstone (Cunningham et al., 2009; Cloete, 2010). However, the subsurface environment is a well-established, diverse and geologically complex domain that comprises a substantial percentage of the Earth's biomass in the form of microorganisms where important biogeochemical cycles work to sustain life (Stevens et al., 1993; Phelps et al., 1994; Fredrickson & Onstott, 1996; Colwell et al., 1997; Onstott et al., 1998; Pedersen et al., 2000; Moser et al., 2003; Kieft et al., 2005; Gihring et al., 2006; Onstott et al., 2009; Borgonie et al., 2011; Ragon et al., 2013; Lau et al., 2014; Rajala et al., 2015), therefore, consideration should be given to the interaction between the sequestered CO2 and the subsurface biome, as well as the geological environments.
Comprehensive studies on the subsurface diversity and geochemistry have revealed that microorganisms from all three domains of life have been found in the South African gold, diamond, and platinum mines (Takai et al., 2001; Moser et al., 2003; Kieft et al., 2005; Gihring et al., 2006; Lin et al., 2006a; Lin et al., 2006b; Onstott et al., 2009; Borgonie et al., 2011; Lau et al., 2014; Magnabosco et al., 2014). Therefore, microorganisms are expected to be present at locations considered for geological storage of CO2. These subsurface researchers have developed extensive sampling techniques to ensure that the microorganisms studied from the deep subsurface are not from contamination through mining or sampling activities, but that they represent the geological setting (Baker et al., 2003; Balkwill et al., 2004; Takai et al., 2001; Borgonie et al., 2011). However, it is possible that the diversity of the subsurface communities can be influenced through mixing
Chapter 3 35
of water types and thus can change over time. Some examples include: 1.) when hydrogen-enriched, ultrabasic subsurface fluids mix with oxygenated surface freshwater (Brazelton et al., 2013), 2.) when sulphate-rich water, found at depths above 300 mbs, mix with methane-rich water, found at depths below 300 mbs (Bomberg et al., 2015), and 3.) when the highly saline hydrothermal fluid mixes with low salinity paleo-meteoric water (Ward et al., 2004; Onstott et al., 2006).
Overall, the chemoautotrophic nature of the subsurface ecosystem increases with depth (Onstott, 2005). This means that the microorganisms in the deep subsurface will be more likely to use CO2 to produce compounds needed for survival such as hydrocarbons, but this depends largely on the geochemistry of the rock and formation water, temperature, and availability of electron donors and acceptors (Onstott, 2005). These chemoautotrophic microorganisms gain their energy through the oxidation of hydrogen (H2), methane (CH4),
2+ 2+ hydrogen sulphide (H2S), ammonia (NH3), ferrous iron (Fe ), and manganese (Mn ). Several groups of microorganisms are capable of producing biomass through chemosynthesis, a process through which these microorganisms use chemical energy to
2- fix CO2 and are dependent on the availability of CO2, sulphate (SO4 ), sulphur (S), ferric
3+ - iron (Fe ), oxygen (O2), or nitrate (NO3 ) as electron acceptors (Jørgensen & Boetius, 2007). These nutrient and chemical energy sources are usually generated through radiolysis, thermogenesis, water-rock interactions or microbial activity and the availability thereof in the subsurface determines the microbial activity in such environments (Kieft et al., 2005; Lin et al., 2006b; Onstott et al., 2006; Etiope & Sherwood Lollar, 2013; Lau et al., 2014). Since these sources are very limited, the activity of the subsurface microorganisms is generally very slow (Phelps et al., 1994; Teske, 2005; Jorgensen & D’Hondt, 2006; Jorgensen & Boetius, 2007; Onstott et al., 2009), but can have both direct and indirect effects. Direct effects include mineral formation/degradation, corrosion, and changes in the porosity due to biofilm formation. Indirect effects include redox and pH changes (Basava- Reddi, 2012), due to interaction with the geological setting.
Although the microbial communities of the deep subsurface are dominated by four anaerobic physiological types - the methanogens, the sulphate/sulphur-reducing bacteria, the fermentative anaerobes, and the Fe(III)-reducing bacteria (Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013), it has been shown that the deep subsurface is an active biosphere that contains a diverse
Chapter 3 36
group of microorganisms with most, if not all, of the physiological types of microorganisms that are thought to be significant in sedimentary environments found in the deep subsurface (Parkes et al., 1994; Lovley & Chapelle, 1995; Whitman et al., 1998; Sahl et al., 2008; Morozova et al., 2010).
3.1.1 Geological Sequestration Sedimentary Rock
During geological carbon sequestration, the CO2 is stored in formations such as sandstone because of the porosity and permeability of the rock (Price & Smith, 2008; Metz et al., 2005). Sandstone (Figure 3.1), also known as arenite, is a clastic sedimentary rock which is composed mainly of sand-sized minerals or rock grains (Pettijohn et al., 1987; Fichter, 2000; Stow, 2005).
Figure 3.1: Sandstone rock with a clastic texture [Taken from Geology4today, (2015)].
Clastic rock consists of fragments of pre-existing calcite, clays, and silicate minerals or rock fragments that were transported by mechanical and chemical weathering and originated through lithification - a process in which sediments compact under pressure, expel connate fluids, and gradually become solid rock. Connate fluids are liquids that were trapped in the pores of sedimentary rocks as they were formed. Rocks, in which sand- sized grains (1/16 mm to 2 mm) are the majority, are called sandstone (Pettijohn et al., 1987; Fichter, 2000; Stow, 2005). The minerals in sandstone, the most common in the Earth’s crust, are quartz, feldspars, and lithic fragments. Matrix may also exist in the interstitial spaces between the rock and mineral grains. Sandstone formations allow
Chapter 3 37
filtration of water and other fluids and are porous enough to store large quantities, making them valuable aquifers (Jackson, 1997; Boggs, 2000).
3.2 AIMS OF THIS CHAPTER
The main aims of this chapter were to: Obtain a study site with similar geological strata and depth as the proposed sites for CCS in South Africa and to do a comprehensive characterization over time of the biogeochemistry of the study site. Analyse the water and gas from the sampling environment for possible changes in the fissure water due to mixing with other water sources, as well as to obtain information regarding the geochemical balance in the borehole’s groundwater that will provide insight into the possible electron donor and acceptor options to sustain biological cycling. Complete extensive diversity studies on the subsurface biome in order to obtain a profile of the different species present, as well as to investigate if any natural shifts in the microbial community structure occurred over time before the introduction of
additional CO2.
Chapter 3 38
3.3 MATERIALS AND METHODS
Unless otherwise specified, the following were applied throughout this research: 1.) All chemicals were obtained from BDH Chemicals Ltd, B&M Scientific cc., BD Biosciences, Bio-Rad Laboratories, Inc., Sigma-Aldrich®, or Merck Millipore. 2.) For media preparations, uniLAB® quality reagents were used and for buffer preparations and all molecular techniques, univAR® quality reagents were used. 3.) All gases were obtained from Air Liquide (Pty) Ltd. 4.) The Nano-pure water was 18 MΩ.cm-1 BarnsteadTM NanopureTM water (Thermo Scientific) that was filtered and UV treated. 5.) The distilled water (dH2O) used was 15 MΩ.cm-1 BarnsteadTM TII water (Thermo Scientific) that was filtered and UV treated. 6.) Sterilization involved autoclaving for 20 minutes at 121°C and 103.4 kPa (1.034 bar). 7.) All graphs were made with Microsoft® Excel (Microsoft® Corporation).
Sampling and analyses of the fissure water, gas, and subsurface biome, to characterize the borehole’s groundwater, included sampling techniques that were designed to maintain subsurface conditions with preservative addition for reliable analyses. Unless otherwise specified, the methods and equipment used for sampling in the mine are standard operating procedures and previously published methods that were developed and adapted over an extensive period of time by members of the Deep Subsurface sampling team (Takai et al., 2001; Lippmann et al., 2003; Moser et al., 2003; Ward et al., 2004; Lin et al., 2006a; Sherwood Lollar et al., 2008; Onstott et al., 2009; Stepanauskas, 2012; Purtschert et al., 2013; Magnabosco et al., 2014). These researchers are part of the DCO Deep Life research program and they are international collaborators, interacting with the Extreme Biochemistry Research group at the University of the Free State (Bloemfontein, RSA), which has been the focal point for this interaction since the year 2000. Samples were transported back to the University of the Free State (UFS) for the necessary processing, while selected samples were sent to the various collaborators for specific analyses.
3.3.1 Sampling Requirements
To understand the deep carbon cycle and its interaction with the subsurface biome, the major physical, chemical and biological conditions for geologic carbon sequestration had to be considered. Deep carbon sequestration would require storage in porous rock at depths of ~800 mbs, an environment with high pressure and temperature (Benson & Cole,
Chapter 3 39
2008; Cloete, 2010; Nondorf et al., 2011). Therefore, this research involved subsurface biomes from geological strata and depth similar to the proposed sites for CCS in South Africa (Cloete, 2010). The ideal site was one where sampling of the microbial community could be conducted >600 mbs in a permeable sandstone region. The Star Diamonds mine made such a site available for at least a 3 year period.
3.3.2 The Study Site
The Star Diamonds mine (Latitude South: 28°19.062’, Longitude East: 026°47.083’, Elevation: 1464 masl) (Figure 3.2) is located approximately 40 km south of Welkom, in the Theunissen district of the Free State province in South Africa and is currently owned by the Frontier Mining Projects (Previously owned by Petra Diamonds).
Figure 3.2: The Star Diamonds mine with shaft 4 that was used as the study site for this research.
The mine is situated east of a dolerite hill on the Adelaide Subgroup of the Beaufort Group (Vermaak, 2012). The first 15 levels inside the mine form part of the Beaufort Group. The Ecca Group is found between levels 15 and 16 with the Dwyka Group from level 16 downwards. The Beaufort Group is the third of the main subdivisions of the Karoo Supergroup of geological strata in South Africa and consists mainly of sandstones and
Chapter 3 40
shale, deposited in the Karoo Basin from the middle Permian to the early part of the middle Triassic periods (Rubidge, 2005; Selden & Nudds, 2011).
3.3.3 The Borehole
The borehole was drilled approximately 0.6 m into the Karoo sandstone until fissure water was intersected (Figure 3.3) and cased (Figure 3.4) to fit the sampling equipment. A diamond drill and compressed air was used to drill the borehole. By not using service water in the drilling process, chances of additional contamination introduced from the service water, were limited. The borehole was drilled on 19 March 2012 and the first sampling occurred within 48 hours, after safety clearance was granted for the site.
Figure 3.3: An approximate position of the borehole, intersecting the Karoo sandstone fissure of the Star Diamonds mine (Supplied by Petra Diamonds/Frontier Mining).
Figure 3.4: The encased borehole, intersecting the Karoo sandstone fissure water, which
was used for sampling in the Star Diamonds mine.
Chapter 3 41
The borehole is situated approximately 640 mbs with a site elevation of 824 metres above sea level (masl) and can be found at shaft 4, level 15 East, cross cut 24. The coordinates of the borehole are: x: -133399.4, y: -19986.8, z: -872.140. This borehole was exclusively used for the research purpose of this project through the courtesy of the team from the Star Diamonds mine (Petra Diamonds/Frontier Mining).
3.3.4 Fissure Water and Gas Sampling and Analyses
Fissure water and gas were sampled through an organic-free manifold (Figure 3.5) that was connected to the casing inside the borehole. This manifold enabled distribution of the water through quick connect clips to the different sampling vials, thus enabling simultaneous sampling of multiple samples, but it also allowed for contamination free sampling by preventing direct contact of the fissure water with the mine air. It also contained a valve for regulation and measurement of the water and gas flow.
Figure 3.5: The organic-free manifold used for sampling, connected to the casing inside the borehole.
Before each sampling event, all manifolds and other sampling equipment were thoroughly cleaned with soapy water and rinsed with Nano-pure water, followed by 70% (v/v) ethanol (EtOH). Equipment was air dried in a laminar flow cabinet (Labotec®), wrapped in autoclave bags and sterilized by autoclaving. Once on-site, the manifold was attached to the borehole and flushed for several minutes with the fissure water by opening all valves before collecting any samples. All samples were collected through sterile Polytetrafluoroethylene (PTFE) tubing (Watson-Marlow) or sterile Masterflex® Norprene
Chapter 3 42
tubing (Cole-Parmer®) that was attached to the quick connect clips of the manifold. All equipment, sampling vials and samples were handled on-site with latex gloves. Unless otherwise specified, fissure water and gas samples were sampled and analysed using previously published methods (Sherwood Lollar et al., 2002; Lippmann et al., 2003; Moser et al., 2003; Ward et al., 2004; Lin et al., 2006a; Onstott et al., 2006; Sherwood-Lollar et al., 2008; Purtschert et al., 2013; Lau et al., 2014).
3.3.4.1 On-Site Physicochemical Analyses
Several on-site analyses were conducted during sampling to monitor the basic geochemistry of the fissure water for indications of any changes or mixing that might occur inside the borehole over time. All these analyses and measurements were done in triplicate according to the procedures of Moser et al., (2003); Lin et al., (2006a); and Lau et al., (2014).
Field parameters (Figure 3.6A) included pH, temperature, salinity, and electrical conductivity (EC), measured with the ExStik®II EC500 handheld probe (EXTECH® Instruments) and oxidation reduction potential (ORP), measured with the ExStik®II RE300 handheld probe (EXTECH® Instruments). All probes were calibrated before sampling using the standard solutions and operating procedures provided by the manufacterer.
Visual colorimetric CHEmets® kits (CHEMetrics Inc.) (Figure 3.6B) were used to measure
2+ the amount of dissolved oxygen (DO), hydrogen sulphide (H2S), dissolved iron (Fe ), total iron, and hydrogen peroxide (H2O2) that is formed from radiolysis of water (H2O) (Chivian et al., 2008). All measurements were conducted according to the manufacturer’s instructions. To eliminate contamination from the casing inside the borehole, all dissolved and total iron samples were filtered using a sterile 0.45 µM cellulose acetate membrane syringe filter (GVS Filter Technology Inc., Lasec) prior to the CHEMets® analyses. For the DO, the funnel provided with the kit was connected to the sampling manifold through a syringe-tipped Masterflex® Norprene tube (Cole-Parmer®) and water was allowed to slowly overflow (Figure 3.6C). Before the ampoule was inserted, care was taken that no bubbles could be observed inside the funnel.
Chapter 3 43
To determine the water flow rate, a 1 L measuring cylinder was filled with water flowing from the borehole (Figure 3.6D) and the time for the cylinder to reach the 1 L mark was recorded. For the gas flow rate, a specifically designed gas stripper (Figure 3.6E) was used (Courtesy of B. Sherwood Lollar) according to the procedure of Ward et al., (2004). The gas stripper was submerged in a bucket that was half filled with fissure water from the borehole. All the taps were opened and the release valve closed. The graduated Perspex beaker was completely filled with fissure water through the telescopic tube. After all the air was displaced from the gas stripper and the rubber tubing, the taps were closed again. The borehole was connected to the inlet tube and the tap was opened to let gas collect inside the graduated Perspex beaker. The time for the gas to displace a fixed volume of water was recorded (Figure 3.6F).
B A C
D E F
Figure 3.6: On-site equipment used for (A) handheld probe measurements, (B) CHEMets®
kits, (C) dissolved oxygen, (D) water flow rate, and (E and F) gas flow rate.
Chapter 3 44
3.3.4.2 Collection of Fissure Water for Geochemical Analyses
For the geochemical analyses, different types of sampling vials were used that had to be prepared before sampling according to the procedures of Moser et al., (2003); Ward et al., (2004); Lin et al., (2006a); and Lau et al., (2014). All sample bottles were prewashed overnight with 10% (v/v) hydrochloric acid (HCl) (Rotzsche, 1991), followed by rinsing three times with Nano-pure water. This was necessary to dissolve and remove all carbonates, iron hydroxides and organic compounds and impurities. All serum vials used were, after being washed and rinsed with the Nano-pure water, allowed to dry before being wrapped in foil and baked overnight in an annealing oven at 450°C. Blue chlorobutyl stoppers (Bellco Glass Inc.) and Teflon septa were washed with 1 M sodium hydroxide (NaOH) and rinsed in Nano-pure water before use. Cleaned serum vials were sealed with the rubber stoppers, crimp sealed with an aluminium cap and autoclaved. For the serum vials that were flushed, a needle was inserted through the chlorobutyl rubber stopper and the vial was pressurized for ten seconds at 50 kPa (0.5 bar) by connecting it directly to the nitrogen gas (99.999%) with a sterile Tygon tube (Thermo Fisher Scientific) and a sterile in-line 0.2 µM IsoporeTM membrane filter (Merck Millipore) that was inserted into a sterile 25 mm in-line Polyacetal Gelman holder (Pall Life Sciences, Sigma-Aldrich®) (Figure 3.7A). A second needle was inserted as an outlet (Figure 3.7B) and the vial was flushed for one minute with nitrogen. The second needle was slowly removed in two movements and the vial was again pressurized for ten seconds, where after all the vials were autoclaved.
Unless otherwise stated, sample bottles were completely filled. For the filtered samples, water was collected on-site through a sterile 0.2 µM cellulose acetate membrane syringe filter (GVS Filter Technology Inc., Lasec) (Figure 3.7C) to remove particulate matter and to reduce the biological activity in the sample. Serum vials were filled on-site by connecting a sterile syringe-tipped Masterflex® Norprene tube (Cole-Parmer®) to the quick connect clip of the sampling manifold. A needle was added onto the syringe tip and inserted through the blue chlorobutyl rubber stopper. A second needle was inserted as an outlet (Figure 3.7D) and water was allowed to fill the vial until the water came out through the outlet needle.
Several preservatives were added to some of the sample vials before sampling took place.
Poisons such as cadmium chloride (CdCl2) and saturated Mercury(II) chloride (HgCl2) were
Chapter 3 45
used to inhibit biological activity after sampling. Acids such as concentrated hydrochloric acid (HCl), sulphuric acid (H2SO4), and nitric acid (HNO3) were used for better preservation of the species being analysed.
The following samples were taken for fissure water chemistry analyses: For anion analyses, a 500 ml Nalgene bottle was filled and measured by ion chromatography (IC)
- − using a DX-120 IC system (Dionex). For formate (HCOO ) and acetate (C2H3O2 ), a 60 ml serum vial was, sealed with a blue chlorobutyl rubber stopper (Bellco Glass Inc.), was filled three quarters by filtering the water and measured by IC using a DX-320 IC system
(Dionex). For cation analyses, five drops of concentrated HNO3 was added to a 50 ml falcon tube and the tube was filled with filtered water. Cations were measured using inductively coupled plasma (ICP), Optima 3000 DV (PerkinElmer Inc.). For the ammonia
+ (NH3) and ammonium (NH4 ) sample, a 40 ml serum vial was sealed with a blue chlorobutyl rubber stopper (Bellco Glass Inc.), 200 µl concentrated H2SO4 was added to decrease the pH of the water sample below five, and autoclaved. The vial was filled with filtered water and the concentrations were measured by the phenol/hypochlorite method (Parsons et al., 1984). The sulphur species sample was collected in the same way as described for formate/acetate, except that the serum vial was also flushed with nitrogen
− (99.999%), before sampling and contained 0.2 g CdCl2. The bisulfide (HS ) content was measured by filtering and dissolving the zinc sulphide (ZnS) precipitate with 0.1 M HCl, and measuring the zinc (Zn) content through inductively coupled plasma atomic emission
2− spectroscopy (ICP-AES), Optima 4300 DV (PerkinElmer Inc.), while the sulphate (SO4 )
2- and thiosulphate (S2O3 ) content of the filtrate was measured by IC using a DX-320 IC system (Dionex). A 500 ml Nalgene bottle was filled three quarters to obtain a
3− phosphorous (P) sample and phosphate (PO4 ) concentrations were measured with a DX- 320 IC system (Dionex). For the total nitrogen (TN), the dissolved organic carbon (DOC), and the dissolved inorganic carbon (DIC), four 250 ml glass bottles with PTFE lined caps (Thermo Fisher Scientific) and eight 125 ml High-density polyethylene (HDPE) bottles (Lasec) were used. The glass bottles were also baked in an annealing oven as previously discussed and all the bottles were filled with filtered water using a Whatman Polycap 36TC, 0.2 µM capsule filter (Thermo Fisher Scientific). The TN was measured simultaneously with the DOC using a TOC-VCSH carbon analyser (Shimadzu) with a TNM-L nitrogen analyser (Shimadzu). The DIC was eliminated through acidification and sparging and the DOC was combusted and measured using a non-dispersive infrared sensor (NDIR). The TN was measured by a chemi-luminescence detector connected in
Chapter 3 46
series with the NDIR. DIC was measured on an Aurora 1030W TOC Analyser (Moser et al., 2003; Lin et al., 2006a; Onstott et al., 2006; Lau et al., 2014).
A B C D
Figure 3.7: Methods used for (A) the sterile 0.2 µM IsoporeTM membrane filter that was
inserted into a sterile 25 mm in-line Polyacetal Gelman holder, (B) flushing serum vials with nitrogen gas, (C) filtering water through a sterile 0.2 µM
syringe filter, and (D) on-site sampling of serum vials.
3.3.4.3 Collection of Gas and Fissure Water for Gas Chemistry and Isotope Analyses
For gas composition, 150 ml serum vials were acid washed, baked and prepared as discussed in Section 3.3.4.2. Saturated HgCl2 (100 µl) was added and the vials were sealed with rubber stoppers and crimp sealed with an aluminium cap. The vials were evacuated by connecting them directly to a vacuum pump (Edwards) with a rubber tube, ending in a syringe tip and a needle that was inserted through the rubber stopper (Figure 3.8A). The pump was switched on until the gauge was at approximately 0.3 mbar and the reading started to slowly drop. When the gauge reached 0.02 mbar, the needle was removed. Once on-site, eight evacuated 150 ml serum vials were filled with gas according to the procedures of Sherwood Lollar et al., (2002) and Ward et al., (2004). Gas samples were obtained by closing the inlet tap from the gas stripper. A needle was connected to the outlet valve and with the tap still open, the needle was inserted into the serum vial for at least two minutes (Figure 3.8B). After the vial was filled, the outlet tap was closed and the needle removed. A 50 ml syringe was connected to the inlet tube and the tap was opened
Chapter 3 47
to fill the syringe with gas. A needle was connected to the syringe and the gas was injected into the serum vial until the plunger pushed out when released. This indicated that the vial was over pressurized. When the gas flow rate was too slow, evacuated 150 ml serum vials were simply filled with fissure water as discussed in Section 3.3.4.2 and again over pressurized by adding additional 20 ml fissure water. While the vials were filled, they were submerged inside a bucket with fissure water (Figure 3.8C). The headspace that formed inside the serum vial was then used as the sample.
Gas compositions that were tested for by gas chromatography (GC) included iso/n-C1-C5, oxygen (O2), and nitrogen (N2) using a thermal conductivity detector, hydrogen (H2) and carbon monoxide (CO) using a reduced gas detector, carbon dioxide (CO2) and methane
(CH4) using a flame ionization detector. Other gases tested for included helium (He), argon
(Ar), nitrous oxide (N2O), and methanethiolate (CH3S). Ultra-high purity (UHP) argon (Ar) was used as the carrier gas, but when measuring Ar, UHP He was used. Sample dilution to instrumental linear response range was conducted using UHP Ar/He.
Additional gas samples that could include noble gas analyses were taken according to the procedure of Lippmann et al., (2003), with modifications from the laboratory of B. Sherwood Lollar. Copper tubes with a 7 mm outer diameter were cut to approximately 70 cm in length. This was done just before sampling to limit the amount of oxidation inside them. The tubes were flushed by connecting them directly to the nitrogen gas (99.999%) with a sterile Tygon tube (Thermo Fisher Scientific) and a sterile in-line 0.2 µM IsoporeTM membrane filter (Merck Millipore) that was inserted into a sterile 25 mm in-line Polyacetal Gelman holder (Pall Life Sciences, Sigma-Aldrich®) (Figure 3.8D). After flushing, the ends were sealed with parafilm (Lasec). Once on-site, four copper tubes were connected in sequence using Tygon tubing. The tubes were pushed at least one centimetre into the Tygon tubing to ensure a tight fitting. At the end of the last copper tube (furthest from the borehole), a longer piece of Tygon tubing was attached and submerged into a beaker of fissure water (Figure 3.8E). The other end of the copper tubes (closest to the borehole), was attached to the outlet valve of the gas stripper discussed in Section 3.3.4.1. The outlet tap was opened and the four tubes were flushed by allowing the gas to flow through for approximately 15 minutes. Flow of gas could be established by ensuring that bubbles were formed at the end that was submerged into the bucket with fissure water. After sampling, each of the four copper tubes was sealed with a crimper at both ends (Figure 3.8F and G) and submerged into a beaker of fissure water to ensure no leaks. Tubes were sealed one
Chapter 3 48
at a time, starting with the copper tube furthest from the borehole and allowing one minute of purging before sealing the next tube.
A B C
D E F G
Figure 3.8: Methods used for (A) evacuating serum vials, (B) collecting gas samples on- site, (C) collecting water samples for gas when the gas flow was too slow, (D) flushing of the copper tubes with nitrogen gas (99.999%), (E) preventing air
from flowing back into the copper tubes, and (F and G) crimp sealing of the
copper tubes.
The concentrations of the dissolved gases were derived from the gas volume abundance, the ratio of water to gas flow rates, and Henry's law constants according to the procedure of Andrews & Wilson, (1987) and are considered to be minimum estimates due to the potential degassing of fluid internally within a partially depressurized fracture zone (Lippmann et al., 2003).
2 13 Isotope analyses of the gas included δ H and δ C for methane (CH4). For the fissure
18 2 water isotope analyses of δ OCH4 and δD/δ HCH4 (deuterium), the sample was collected in a 60 ml narrow mouth Nalgene bottle (Moser et al., 2003; Lin et al., 2006a; Onstott et al.,
14 2006; Lau et al., 2014). For the δ CDIC, a 500 ml Nalgene bottle, containing 1 ml of a saturated HgCl2 solution, were filled. Immediately after sampling, the bottle was sealed to
14 prevent contamination by the CO2 in air. The conventional Radiocarbon Age and δ C are reported according to the procedures of Stuiver & Polach, (1977). The fraction modern (F)
Chapter 3 49
is the blank corrected fraction modern normalized to δ13C of -25 g.L-1 (Donahue et al., 1990). The δ13C normalization was performed using δ13C measured by accelerator mass spectrometry (AMS), thus accounting for AMS fractionation. The reported errors comprise statistical errors in sample and standard determinations, combined in quadrature with a system error based on the analyses of an on-going series of measurements on an oxalic acid standard.
3.3.5 Sandstone Sampling and Analyses
Karoo sandstone (Figure 3.9) was obtained from the Star Diamonds mine to be used in several different experiments. The sandstone was collected from the same area where the borehole is located.
Figure 3.9: The Karoo sandstone obtained from the Star Diamonds mine.
3.3.5.1 Crushing and Sterilizing
The sandstone was crushed using a jaw crusher and separated according to size using a 2.8 mm and a 4.7 mm strainer (4.7 mm < X > 2.8 mm) (Figure 3.10).
Figure 3.10: Separation of the crushed rock using a 2.8 mm and a 4.7 mm strainer.
Chapter 3 50
The crushed rock was sterilized through either autoclaving, acid washing, or both. Acid washing was conducted by an overnight wash step in 10% (v/v) HCl (Rotzsche, 1991), followed by rinsing three times with sterile Nano-pure water. This was all done under sterile conditions in a laminar flow cabinet (Labotec®). When both methods were used, the rock was first washed and rinsed before being autoclaved.
3.3.5.2 X-Ray Fluorescence Analyses
The composition of the sandstone was analysed by X-Ray fluorescence analyses (XRF) (Rollinson, 1993; Von Eynatten et al., 2003) for any possible changes that might have occurred during processing. XRF analyses were conducted at the Department of Geology at the UFS (Bloemfontein, RSA).
The crushed sandstone was milled to powder using a tungsten mill, where after 8 g of the powder was added to 3 g of Hoechst wax (C6H8O3N2). This was mixed for 20 minutes in a Turbula mixer to ensure that the sample was mixed homogeneously and pressed to pressures greater than 395 N.m-1 to make a pellet for measuring the trace-elements and sodium. The total loss on ignition (LOI) was calculated by heating the powdered sample to 110°C (dehydration) for 2 hours and then over night at 1050°C. A flux of 0.2445 g lanthanum oxide (La2O3), 0.705 g lithium tetraborate (Li2B4O7), 0.5505 g lithium carbonate
(Li2CO3), and 0.02 g sodium nitrate (NaNO3) was added to 0.28 g of the powdered sample. This mixture was then heated in a platinum (Pt) crucible to 1000°C for approximately 5 minutes until a consistent fluid formed. The fluid was poured into a mould and pressed to create a fusion disc for measuring the major-elements. Analyses were conducted by Xray fluorescence spectrometry using an Axios XRF spectrometer (PANalytical) with a rhodium (Rh) tube. Major-element oxides were recalculated to 100% with the LOI included.
3.3.6 Diversity Analyses of the Deep Subsurface Microbial Biome
The number of species (the species richness) and the sizes of the species populations (the species evenness) within a community are both important parameters to consider when describing the diversity within a community (Liu et al., 1997). However, the portion of microbial species that can be cultured and identified based on their physiological and biochemical properties through the use of traditional culture-dependant methods only
Chapter 3 51
represent a small fraction of the existing diversity (Amann et al., 1995; Liu et al., 1997). It is also likely that the diversity within a community will change if the culture conditions differ even slightly from the conditions of their original environment. This will introduce essential biases in the diversity of the community due to the methods used for isolation (Dunbar et al., 1997; Liu et al., 1997).
Therefore, for a more comprehensive characterization of the subsurface biome, samples were collected for genomic deoxyribonucleic acid (gDNA) and total ribonucleic acid (RNA) isolations by using various types of filters and larger quantities of fissure water. Unless otherwise specified, subsurface microbial biomes were sampled and analysed according to the procedures of Takai et al., (2001); Moser et al., (2003); Lin et al., (2006a); Lau et al., (2014); and Magnabosco et al., (2014).
3.3.6.1 Cornelius® Canister Sampling
For continual monitoring of the diversity over time, fissure water was collected anaerobically in a 12.6 L stainless steel Cornelius® canister (Cornelius® Inc.) (Figure 3.11) according to the procedures of Takai et al., (2001); Moser et al., (2003); and Lin et al., (2006a). Before each sampling, the canister was thoroughly cleaned with soapy water and rinsed with Nano-pure water, followed by 70% (v/v) EtOH. The canister was air dried in a laminar flow cabinet (Labotec®) and sterilized together with all tubing and fittings needed to connect the canister to the sampling manifold attached to the borehole. The outlet valve on the cap was opened before autoclaving (Figure 3.11A). After sterilization, the outlet valve was closed and the canister was allowed to cool to room temperature. The canister was flushed for 10 minutes at 50 kPa (0.5 bar) by connecting it directly to the nitrogen gas (99.999%) with an autoclaved inlet adaptor fitted to sterile Tygon tubing (Thermo Fisher Scientific) and a sterile in-line 0.2 µM IsoporeTM membrane filter (Merck Millipore) that was inserted into a sterile 25 mm in-line Polyacetal Gelman holder (Pall Life Sciences, Sigma- Aldrich®) (Figure 3.11B). During flushing the outlet valve was opened. Before canisters were used, the outlet valve was quickly opened to ensure that the canister maintained positive pressure.
On-site, the inlet of the canister was directly connected to a quick connect clip of the sampling manifold by using sterile PTFE tubing (Watson-Marlow) (Figure 3.11C). As water
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flowed into the canister, the outlet valve was opened until the canister was overflowing (Figure 3.11D). In the laboratory, the fissure water was displaced from the canister through the addition of nitrogen gas (99.999%) at 50 kPa by connecting the inlet of the canister directly to the nitrogen with an autoclaved inlet adaptor fitted to sterile Tygon tubing (Thermo Fisher Scientific) and an in-line 0.2 µM IsoporeTM membrane filter (Merck Millipore) (Figure 3.11E). The fissure water is displaced using positive pressure (Figure 3.11F) and the cells were collected on sterile Supor® 200 membrane disc filters (Pall Life Sciences) that were inserted into sterile 47 mm in-line Polycarbonate Gelman holders (Pall Life Sciences, Sigma-Aldrich®) (Figure 3.11G). After filtration, the filters were stored in cryogenic vials (Thermo Fisher Scientific) at -20°C until further processing.
A
D B C
E F G
Figure 3.11: Methods used for (A) sterilizing the canister with the outlet valve opened, (B) flushing the canister, (C and D) collecting fissure water on-site, and (E-G)
filtering water from the canister.
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3.3.6.2 SterivexTM Filter Sampling
As an alternative means to the Cornelius® canisters, sterile SterivexTM - GP 0.22 µM, polyethersulone (PES) filters (Millipore Corporation) (Figure 3.12A) were connected to the borehole to collect biomass according to the procedure of Brazelton et al., (2012). The filter caps were removed and the filters were attached to the quick connect clips of the sampling manifold by using a sterile syringe-tipped Masterflex® Norprene tube (Cole- Parmer®) (Figure 3.12B). The filters had to stay connected inside the mine for at least one month to obtain larger quantities of biomass. Filters were stored at -20°C until further processing.
A B
Figure 3.12: (A) The SterivexTM filters used to (B) collect biofilm on-site through direct connection to a quick connect clip of the sampling manifold.
3.3.6.3 Tangential Flow Filtration
In order to obtain a concentrated sample of the subsurface biome, large volumes of fissure water were filtered and concentrated using tangential flow filtration (TFF) according to the procedure of Kuwabara and Harvey, (1990). Fissure water was collected in the mine with sterile 50 L Nalgene Carboy bottles. The water was transported back to the laboratory where TFF was carried out under sterile conditions inside a laminar flow cabinet (Labotec®) (Figure 3.13A). The water was filtered using a sterile 100 000 NMWC pore size, polysulfone (PS), ultrafiltration hollow fibre cartridge [63.5 cm (length) x 5.1 cm (outer diameter)] (GE Healthcare).
Chapter 3 54
This process was repeated on-site to obtain sufficient biome quantities (Figure 3.13B). A 50 L sterile Nalgene Carboy bottle was also filled with un-concentrated fissure water, and together with the concentrated water, transported back to the laboratory where it was stored at 4°C to be used in different experiments as needed.
A B
Figure 3.13: Tangential flow filtration of the fissure water (A) inside the laboratory and (B) inside the mine.
3.3.6.4 Massive Filter Sampling
For collecting biomass from larger quantities of fissure water, a massive filter (Figure 3.14A) was used according to the procedures of Lau et al., (2014) and Magnabosco et al., (2014). Before each sampling, the filter holder was thoroughly cleaned with soapy water and rinsed with Nano-pure water, followed by 70% (v/v) EtOH and air dried in a laminar flow cabinet (Labotec®). A 0.1 µM Osmonics Nylon filter cartridge (GE Healthcare) was inserted into the filter holder and the lid was closed. The massive filter, together with all tubing and fittings used, were placed in an autoclave bag and sterilized before use.
On-site, the inlet of the massive filter was directly connected to a quick connect clip of the sampling manifold using sterile PTFE tubing (Watson-Marlow) (Figure 3.14B) and the outlet of the massive filter was directly connected to a flow meter (Figure 3.14C). The filter had to stay connected inside the mine for at least one month to obtain large quantities of biomass.
Chapter 3 55
A B C
Figure 3.14: (A) A massive filter used for collecting biofilm from fissure water in larger
quantities through (B) direct connection to a quick connect clip of the sampling
manifold to the inlet side of the filter and (C) the flow accumulator connected to the outlet side of the filter.
Unfortunately the pressure from the borehole was not enough to obtain a flow of water through the filters and the filters had to be removed after three days. A new massive filter was prepared as discussed previously, but this time connected to a peristaltic pump (Millipore Corporation) and a Nalgene Carboy bottle that served as a reservoir once the bottle was filled (Figure 3.15). The cap of the Carboy bottle had three connections – one inlet and two outlets (Figure 3.15A). The inlet of the cap was directly connected to a quick connect clip of the sampling manifold using sterile PTFE tubing (Watson-Marlow), as shown before in Figure 3.14B. The first outlet was connected to the peristaltic pump that was set to operate at maximum speed (600 rpm) with sterile Masterflex® Norprene tubing (Cole-Parmer®) (Figure 3.15B) and the second outlet was left open as an overflow (Figure 3.15C). A sterile 0.2 µM nylon filter cloth was placed underneath the cap to cover the opening of the Carboy bottle. Both the inlet of the cap and the outlet that was connected to the peristaltic pump had a tube connected on the inside of the cap (Figure 3.15D) that went through the filter all the way to the bottom of the Carboy. This was done so that the biofilm was collected inside the carboy and not on the filter. The filter was replaced every second day. The peristaltic pump was connected to the inlet of the massive filter (Figure 3.15E) and the outlet of the massive filter was directly connected to a flow accumulator, as shown before in Figure 3.14C. The numbers on the flow accumulator were recorded before being attached to the filter. This time enough pressure was created for the water to keep on flowing through the filter which was left connected to the borehole for one month.
Chapter 3 56
Upon retrieval of the massive filter, immediately after disconnecting the filter from the borehole, the water inside the filter holder was discarded through the inlet/outlet and the filter holder was filled with a sterile, salt-saturated RNA preservative solution [20 mM ethylenediaminetetraacetic acid (EDTA), 0.3 M sodium citrate, 4.3 M ammonium sulphate, pH 5.2, adjusted with concentrated sulphuric acid (H2SO4)] (Lau et al., 2014). The filter holder was sealed with sterile threaded plugs, transported back to the laboratory at the UFS (Bloemfontein, RSA), and stored overnight at 4°C to ensure saturation of all the membrane layers of the filter. The filter was then removed from the filter holder under sterile conditions, wrapped in aluminium foil and stored at -80°C until further processing.
The Nalgene Carboy that contained large amounts of cells, was closed with a sterile cap and the water was transported back to the laboratory at the UFS (Bloemfontein, RSA) and stored at 4°C to be used in different experiments as needed.
A B
C D E
Figure 3.15: (A) The cap of the Carboy bottle with one inlet and two outlets with (B) one of the outlets connected to the pump, and (C) the other outlet left open as an overflow, (D) the tube connected on the inside of the cap, and (E) the pump
connected to the inlet of the massive filter.
Chapter 3 57
3.3.6.5 Polyvinyl Chloride Cartridge Sampling
As an alternative to the massive filters, sterile Polyvinyl chloride (PVC) cartridges, filled with crushed sandstone (Figure 3.16) were connected to the borehole (Moser et al., 2003). These cartridges were custom-made by the Department of Instrumentation at the UFS (Bloemfontein, RSA). The cartridge consisted of a hollow PVC casing that was fitted on the one side with a quick connect clip and a reducer on the other side. Each cartridge also had two screw caps for proper sealing during storage (Figure 3.16A). The sandstone was added to provide surfaces for microbial attachment and colonization. A quick connect clip and reducer was attached to the cartridge and together with the screw caps, wrapped in aluminium foil and sterilized.
On-site the cartridges were directly connected to the sampling manifold (Figure 3.16B) and the cartridges stayed connected to the borehole for one month. Once the cartridges were removed, they were sealed with the sterile caps, wrapped in aluminium foil and transported back to the laboratory at the UFS (Bloemfontein, RSA) where they were stored at -80°C until further processing.
A B
Figure 3.16: (A) The custom made PVC cartridges with its fitting and caps and (B) the
cartridges directly connected to the sampling manifold on the borehole.
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3.3.6.6 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis
Denaturing gradient gel electrophoresis (DGGE) was used to determine the genetic diversity of the microbial communities from the subsurface biome.
3.3.6.6.1 Genomic DNA Isolation
Genomic DNA was isolated from all the filters of the Cornelius® canisters, the SterivexTM filters, as well as from the biome that was concentrated using TFF. The filters were allowed to thaw and cut into small pieces under sterile conditions. The filter pieces, as well as the concentrated TFF water were added to the tubes containing the beads and the gDNA was isolated using the NucleoSpin® Soil kit (Macherey-Nagel) according to the manufacturer’s instructions. The integrity of the gDNA was evaluated using agarose gel electrophoresis. All of the agarose gels used throughout this research consisted of 1% (w/v) agarose in Tris-Acetate-EDTA (TAE) buffer (40 mM Tris, 40 mM acetic acid, 1 mM EDTA, pH 7.4) with 0.6 µg.ml-1 ethidium bromide (EtBr) (Merck Millipore). Genomic DNA (10 µl) was added to 2 µl loading dye (Fermentas) before being separated in the gel at 90 V for 60 minutes. Either MassRulerTM DNA Ladder Mix or O’GeneRulerTM DNA Ladder Mix (Both from Thermo Scientific) was used to determine the size of the amplicons and the gel was visualized using the ChemiDoc XRS UV/Vis Gel Documentation system with Quantity One V.4.5 software (Bio-Rad Laboratories, Inc.). The concentration and purity of the gDNA was measured using the NanoDrop® ND-1000 UV/Vis Spectrophotometer (Inqaba BiotechTM).
3.3.6.6.2 Polymerase Chain Reaction
A polymerase chain reaction (PCR) was conducted on the gDNA to amplify the Archaeal 16S (~600 bp), Bacterial 16S (~1500 bp) and Eukaryal 18S (~1700 bp) rRNA gene fragments using the universal oligonucleotide primers for each gene as indicated in Table 3.1. Control reactions were prepared for each of the different groups using sterile water as the negative control and gDNA from Halobacterium salinarum, Escherichia coli and Saccharomyces cerevisiae as the templates for the positive controls for Archaea, Bacteria and Eukarya, respectively.
Chapter 3 59
Table 3.1: The universal oligonucleotide primers used for amplification of the Archaeal 16S, Bacterial 16S and Eukaryal 18S rRNA gene fragments.
Primer Nucleotide sequence Tm (°C) rRNA gene Reference
Arc333F 5'-TCC AGG CCC TAC GGG-3' 56.9 Archaea 16S Lepp et al., 2004
Arc958R 5'-YCC GGC GTT GAM TCC AAT T-3' 56.6 Archaea 16S De Long, 1992
Bac27F 5'-AGA GTT TGA TCM TGG CTC AG-3' 53.2 Bacteria 16S Turner et al., 1999
Bac1492R 5'-GGT TAC CTT GTT ACG ACT T-3' 49.4 Bacteria 16S Turner et al., 1999
EukA 5’-AAC CTG GTT GAT CCT GCC AGT-3’ 58.9 Eukarya 18S Diez et al., 2001
EukB 5’-TGA TCC TTC TGC AGG TTC ACC TAC-3’ 58.4 Eukarya 18S Diez et al., 2001
The PCR was performed in a total reaction volume of 25 µl using a 2720 Thermal Cycler (Applied Biosystems®). The reaction mixture consisted of 12.5 µl GoTaq® Green master mix (2X) [a premixed ready-to-use solution that contains the Taq DNA polymerase, dNTPs
(400 µM), magnesium chloride (MgCl2) (3 mM) and reaction buffers (pH 8.5)], 1 µl each of both the forward and reverse primers (10 µM), 50 ng.µl-1 template and sterile Nano-pure water filled to the reaction volume.
Optimized reaction conditions for Archaeal 16S consisted of an initial denaturing step at 94°C for 2 minutes, followed by 30 cycles of denaturing at 95°C (30 seconds), annealing at 61°C (30 seconds) and elongation at 72°C (1.5 minutes). A final elongation step of 1 minute at 72°C was added to ensure complete elongation of the amplified product. For Bacterial 16S, optimized reaction conditions consisted of an initial denaturing step at 95°C for 5 minutes, followed by 30 cycles of denaturing at 95°C (30 seconds), annealing at 49°C (45 seconds) and elongation at 72°C (1 minute), with a final elongation step of 10 minutes at 72°C. For Eukaryal 18S, optimized reaction conditions consisted of an initial denaturing step at 94°C for 1.5 minutes, followed by 30 cycles of denaturing at 94°C (30 seconds), annealing at 60°C (1 minute) and elongation at 72°C (45 seconds), with a final elongation step of 5 minutes at 72°C.
The PCR products were visualized on a 1% (w/v) agarose gel. Purification of the PCR products was achieved by excising the desired band from the agarose gel using a non-UV Dark ReaderTM DR-45M transilluminator (Clare Chemical Research) and extracted using the BioFlux BioSpin Gel Extraction Kit (Bioer Technology Co., Ltd) according to the manufacturer’s instructions. The concentration and purity of the purified products were
Chapter 3 60
measured using the NanoDrop® ND-1000 UV/Vis Spectrophotometer (Inqaba BiotechTM). All PCR reactions were done in triplicate and combined into one tube to increase concentrations before purifications were done.
3.3.6.6.3 Nested Polymerase Chain Reaction
A nested DGGE PCR was conducted on all the purified samples to amplify the V3/V4 hypervariable regions for Archaea (~600 bp), Bacteria (~600 bp) and Eukarya (~600 bp) using the DGGE oligonucleotide primers for each gene as indicated in Table 3.2. A GC- clamp was added to the forward primers.
Table 3.2: The DGGE oligonucleotide primers used for amplification of the V3/V4 hypervariable regions for Archaea, Bacteria and Eukarya, respectively. Underlined sequences indicate the GC-clamp.
Primer Nucleotide sequence Tm (°C) V3/V4 Reference
5'-CGC CCG CCG CGC GCG GCG GGC Arc344F-GC GGG GCG GGG GCA CGG GGG GAC 86.2 Archaea Casamayor et al., 2002 GGG GYG CAG CAG GCG CGA-3'
Arc934R 5'-GTG CTC CCC CGC CAA TTC CT-3' 62.9 Archaea Casamayor et al., 2002
5'-CGC CCG CCG CGC GCG GCG GGC Bac341F-GC GGG GCG GGG GCA CGG GGG GCC 85.1 Bacteria Muyzer et al., 1993 TAC GGG AGG CAG CAG-3'
Bac908R 5'-CCG TCA ATT CMT TTG AGT TT-3' 49.9 Bacteria Turner et al., 1999
5’-CGC CCG CCG CGC GCG GCG GGC Euk1AF-GC GGG GCG GGG GCA CGG GGG GCT 84.1 Eukarya Diez et al., 2001 GGT TGA TCC TGC CAG-3’
Euk516R 5’-ACC AGA CTT GCC CTC C-3’ 54.3 Eukarya Diez et al., 2001
The DGGE PCR was performed in a total reaction volume of 25 µl using a 2720 Thermal Cycler (Applied Biosystems®) as discussed in Section 3.3.6.6.2. Optimized reaction conditions for the Archaeal V3/V4 region consisted of an initial denaturing step at 94°C for 2 minutes, followed by 30 cycles of denaturing at 95°C (30 seconds), annealing at 61°C (30 seconds) and elongation at 72°C (1.5 minutes). A final elongation step of 1 minute at 72°C was added to ensure complete elongation of the amplified product. For the Bacterial V3/V4 region, optimized reaction conditions consisted of an initial denaturing step at 95°C for 4 minutes, followed by 30 cycles of denaturing at 95°C (1 minute), annealing at 55°C (1
Chapter 3 61
minute) and elongation at 72°C (1.5 minutes), with a final elongation step of 10 minutes at 72°C. For the Eukaryal V3/V4 region, optimized reaction conditions consisted of an initial denaturing step at 95°C for 4 minutes, followed by 30 cycles of denaturing at 95°C (1 minute), annealing at 56°C (1 minute) and elongation at 72°C (1.5 minutes), with a final elongation step of 10 minutes at 72°C. The DGGE PCR products were visualized on a 1% (w/v) agarose gel.
3.3.6.6.4 Denaturing Gradient Gel Electrophoresis
The products from the nested DGGE PCR were used to construct preliminary diversity profiles using the DGGE DcodeTM universal mutation detection system (Bio-Rad Laboratories, Inc.) as described by Diez et al., (2001). The gradient of urea/formamide denaturants for Archaea and Bacteria ranged from 40% - 60% and for Eukarya ranged from 40% - 65%. Electrophoresis was performed with a 7% (w/v) polyacrylamide gel (acrylamide/bisacrylamide 40%, 37.5:1 ratio, Merck Millipore) in Tris-Acetate-EDTA (TAE) buffer (40 mM Tris, 40 mM acetic acid, 1 mM EDTA, pH 7.4) at 60°C. The entire nested DGGE PCR product was used and separated in the gel at 100 V for 16 hours. The gel was stained with 0.6 µg.ml-1 ethidium bromide (EtBr) (Merck Millipore) for 30 minutes in TAE buffer and visualized using the ChemiDoc XRS UV/Vis Gel Documentation system with Quantity One V.4.5 software (Bio-Rad Laboratories, Inc.).
3.3.6.6.5 Gel Extraction, Re-amplification and Purification
Selected DGGE bands were excised from the gel using the ChemiDoc XRS UV/Vis Gel Documentation system with Quantity One V.4.5 software (Bio-Rad Laboratories, Inc.) and eluted overnight at 50°C in 50 µl sterile Nano-pure water. The concentration and purity of the eluted DGGE products were measured using the NanoDrop® ND-1000 UV/Vis Spectrophotometer (Inqaba BiotechTM).
The concentrations of the recovered DNA were increased through re-amplification of the V3/V4 hypervariable regions from the eluted DGGE products for Archaea (~600 bp), Bacteria (~600 bp) and Eukarya (~500 bp) using the oligonucleotide primers for each gene as indicated in Table 3.3.
Chapter 3 62
Table 3.3: The oligonucleotide primers used for the re-amplification of the V3/V4 hypervariable regions for Archaea, Bacteria and Eukarya, respectively.
Primer Nucleotide sequence Tm (°C) V3/V4 Reference
Arc344F 5'-ACG GGG YGC AGC AGG CGC GA-3' 72.9 Archaea Casamayor et al., 2002
Arc934R 5'-GTG CTC CCC CGC CAA TTC CT-3' 62.9 Archaea Casamayor et al., 2002
Bac341F 5'-CCT ACG GGA GGC AGC AG-3' 58.9 Bacteria Muyzer et al., 1993
Bac908R 5'-CCG TCA ATT CMT TTG AGT TT-3' 49.9 Bacteria Turner et al., 1999
Euk1AF 5’-CTG GTT GAT CCT GCC AG-3’ 52.8 Eukarya Diez et al., 2001
Euk516R 5’-ACC AGA CTT GCC CTC C-3’ 54.3 Eukarya Diez et al., 2001
The PCR was performed in a total reaction volume of 25 µl using a 2720 Thermal Cycler (Applied Biosystems®) as discussed in Section 3.3.6.6.2.
Optimized reaction conditions for the Archaeal V3/V4 region consisted of an initial denaturing step at 94°C for 2 minutes, followed by 30 cycles of denaturing at 95°C (30 seconds), annealing at 59°C (30 seconds) and elongation at 72°C (1.5 minutes). A final elongation step of 1 minute at 72°C was added to ensure complete elongation of the amplified product. For the Bacterial V3/V4 region, optimized reaction conditions consisted of an initial denaturing step at 95°C for 4 minutes, followed by 30 cycles of denaturing at 95°C (1 minute), annealing at 53°C (1 minute) and elongation at 72°C (1.5 minutes), with a final elongation step of 10 minutes at 72°C. For the Eukaryal V3/V4 region, optimized reaction conditions consisted of an initial denaturing step at 95°C for 4 minutes, followed by 30 cycles of denaturing at 95°C (1 minute), annealing at 54°C (1 minute) and elongation at 72°C (1.5 minutes), with a final elongation step of 10 minutes at 72°C. The re-amplified PCR products were visualized on a 1% (w/v) agarose gel and purified (Section 3.3.6.6.2).
3.3.6.6.6 Sanger Sequencing and Analyses
The purified DGGE products were sequenced to determine the genetic diversity of the subsurface microbial community. Sanger sequencing was conducted using the ABI Prism® Big DyeTM Terminator Cycle Sequencing Ready Reaction Kit V.3.1 (Applied Biosystems) according to the manufacturer’s instructions.
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The reaction was performed in a total volume of 10 µl and consisted of 0.5 µl premix, 2 µl dilution buffer, 1 µl primer (3.2 pmol.µl-1), template (200-500 ng.µl-1) and sterile Nano-pure water filled to the reaction volume. Reaction conditions consisted of an initial denaturing step at 96°C for 1 minute, followed by 25 cycles of denaturing at 96°C (10 seconds), annealing at 50°C (5 seconds) and elongation at 60°C (4 minutes). The primers in Section 3.3.6.6.5, Table 3.3 were used, and separate reactions were made for the forward and reverse primers.
The PCR products were purified using an EDTA/EtOH precipitation protocol for sequencing clean-up. The sequencing reaction volume was adjusted to 20 µl, transferred to a 1.5 ml eppendorf tube containing 5 µl EDTA and 60 µl absolute EtOH (Merck Millipore), mixed for 5 seconds and allowed to precipitate for 15 minutes at room temperature. The mixture was centrifuged (20 000 x g, 10 minutes, 4°C) using an AllegraTM 25R centrifuge (Beckman CoulterTM) and the supernatant was removed through aspiration, without disturbing the pellet. 70% (v/v) EtOH (60 µl) was added to the pellet and centrifuged once more (20 000 x g, 5 minutes, 4°C). The supernatant was removed through aspiration and the pellet was dried for 5 minutes using a Concentrator 5301 (Eppendorf). The samples were covered in foil and stored in the dark at 4°C until sequenced. Sequencing reactions were conducted with a 3130xl Genetic Analyzer, HITACHI (Applied Biosystems) at the UFS (Bloemfontein, RSA).
For the 16S and 18S rRNA gene sequences, the sequence data was analysed and assembled to create contigs using the Geneious software (V.7.1.7). Analysed sequences for 16S were aligned against the life tree project (LTP) 119 rRNA database from SILVA (Pruesse et al., 2007) using a variant (BLASTN) algorithm (Zhang et al., 2000) of the basic local alignment search tool (BLAST). Analysed sequences for 18S were aligned against the Nucleotide collection (nr/nt) database from the National Centre for Biotechnology Information (NCBI) using the BLASTN algorithm (Altschul et al., 1997), by excluding uncultured and environmental samples in order to obtain the closest family. The first fifteen hits were used to assign higher order classification and were filtered based on 50% identity.
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3.3.6.7 Microbial Diversity Assessments using Targeted 16S and 18S rRNA Gene Sequencing
3.3.6.7.1 Genomic DNA Isolation
Genomic DNA was isolated from the massive filters as well as the PVC cartridges. The massive filters were prepared by removing the filters from the -80°C freezer and allowed to thaw. The filter was cut into ~2 cm sections and the different layers of the filter were separated into the inner fibrous membrane, the massive filter, and the outer fibrous membrane. The different layers were each cut into smaller pieces and used for gDNA isolations by adding the filter pieces to the tubes containing the beads. For the PVC cartridges, the cartridges were removed from the -80°C freezer and allowed to thaw. The rock was removed from the cartridge and added to a tube without beads. The rock was used to break the cells in this case.
The gDNA was isolated using two different methods. The first method used was the NucleoSpin® Soil kit (Macherey-Nagel) according to the manufacturer’s instructions. The second method used was according to the procedures of Lau et al., (2014). The reaction tubes were kept on ice throughout the extraction process to limit nucleic acid degradation. The cells were lysed by adding sterile cetyl trimethylammonium bromide (CTAB) lysis buffer (2X) [8.18 g.L-1 NaCl, 0.5 M EDTA (4 ml), 1 M Tris, pH 8.0 (10 ml), 2 g.L-1 CTAB in
-1 -1 dH2O] containing lysozyme (5 mg.mL ) and proteinase K (0.2 mg.mL ) to the tubes containing the material from the filters. The tubes were vortexed for one minute, incubated in a water bath at 60°C for 30 minutes and rinsed with sterile Tris-EDTA (TE) buffer (1X) [1
M Tris, pH 8.0 (10 ml), 0.5 M EDTA (2 ml) in dH2O]. A phenol/chloroform/isoamyl alcohol (25:24:1 v/v/v) extraction was conducted by adding 4 ml of the phenol/chloroform/isoamyl alcohol to each 5 ml of the lysate. The mixture was incubated in a water bath at 60°C for one minute, added to an ice-bath for 5 minutes and centrifuged (4 300 x g, 10 minutes, room temperature) using an AllegraTM 25R centrifuge (Beckman CoulterTM). The gDNA was precipitated by adding isopropanol (1:1 v/v) to the mixture and gently inverted to mix. The tubes were incubated in an ice-bath for 30 minutes and centrifuged (4 300 x g, 15 minutes, room temperature) using an AllegraTM 25R centrifuge (Beckman CoulterTM). The supernatant was discarded and the pellets were rinsed with ice-cold 75% EtOH and allowed to air dry. Once dried, the pellets were re-suspended in 300 µl sterile TE buffer (1X), containing RNase A (10 µg.ml-1) (Fermentas) and incubated in a water bath at 37°C
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for 30 minutes. NaCl (0.1 M) and two volumes of absolute EtOH were added to the mixture, incubated at -20°C for 30 minutes and centrifuged (11 500 x g, 30 minutes, room temperature) using an AllegraTM 25R centrifuge (Beckman CoulterTM) to collect the gDNA. The gDNA was subjected to quality control as discussed in Section 3.3.6.6.1.
3.3.6.7.2 MiSeq Illumina® Sequencing and Analyses
16S rRNA metagenome sequence analyses were conducted on the gDNA using the MiSeq Illumina® platform. The main steps included performing a standard 16S rRNA gene PCR using 16S forward and reverse primers which contained Illumina® adaptors and target specific sequences. Amplicons were quantified using the NanoDrop ND-3300 UV/Vis Spectrophotometer (Inqaba BiotechTM) to determine the amount needed for the library preparation. An Index PCR-limited cycle PCR was conducted using the Nextera XT DNA Sample Preparation and the Nextera XT Index Kits (Illumina®). Barcodes/indices were added to the 5’ and 3’ ends of the amplicons to enable demultiplexing of the forward and reverse reads. Sequencing reactions were conducted using the Miseq Reagents Cartridge and Incorporation buffer on the Illumina® MiSeq instrument at Inqaba BiotechTM (Hatfield, RSA). The primers and standards used were as recommended by the manufacturer. The data was analysed with the automated analysis platform Metagenomics RAST (MG-RAST) using the default settings (Meyer et al., 2008).
3.3.6.7.3 Roche GS Junior Sequencing and Analyses
18S rRNA metagenome sequence analyses were conducted on the gDNA using the Roche GS Junior platform. The main steps included performing a standard 18S rRNA gene PCR using 18S barcoded forward primers, containing 454 adaptor-tag/barcode- target specific sequences and an 18S reverse primer, containing 454 adaptor-target specific sequences. Amplicons were quantified using the NanoDrop ND-3300 UV/Vis Spectrophotometer (Inqaba BiotechTM) to determine the number of copies (amplicons) per bead for emulsion PCR (emPCR). For the emPCR, the GS Junior Titanium emPCR Reagent Kit (Roche Life Science) was used according to the manufacturer’s instructions. The emPCR product recovery was followed by enrichment of DNA carrying beads and counting thereof to load the PicoTiterPlate (PTP) using the GS Junior Titanium PicoTiterPlate Kit (Roche Life Science). These were sequenced using the GS Junior
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Titanium Sequencing Kit (Roche Life Science) according to the manufacturer’s instructions. Sequencing reactions were conducted with the Roche 454 GS Junior (Roche Life Science) at Inqaba BiotechTM (Hatfield, RSA). The primers and standards used were as recommended by the manufacturer. The data was analysed with MG-RAST using the default settings (Meyer et al., 2008).
3.3.6.7.4 Ion Torrent PGMTM Sequencing and Analyses
Both 16S and 18S rRNA metagenome sequence analyses were conducted on the gDNA using the Ion Torrent Personal Genome MachineTM (Ion PGMTM) platform (Life TechnologiesTM). The 18S library was constructed from the gDNA for up to 400 base-read sequencing using the Ion XpressTM Plus Fragment Library Kit (Life TechnologiesTM) according to the manufacturer’s instructions for enzyme shearing. The 16S library was constructed from the gDNA for a 200 – 400 base-read sequencing using the Ion 16STM Metagenomics Kit (Thermo Fisher Scientific) according to the manufacturer’s instructions. The template was prepared from the 16S library using the Ion PGMTM Template OT2 400 Kit (Life TechnologiesTM) according to the manufacturer’s instructions and sequenced using the Ion PGMTM Hi-QTM Sequencing Kit (Life TechnologiesTM) according to the manufacturer’s instructions. All assessments of the DNA were done using a 2100 Bioanalyzer® (Agilent® Technologies Inc.). The entire 18S region was sequenced and assembled as contigs. For the 16S, the V2-V4, and V6-V9 regions were sequenced. Sequencing reactions were conducted with the Ion Torrent PGMTM (Life TechnologiesTM) at the Central Analytical Facilities (CAF), Stellenbosch University (Stellenbosch, RSA). The primers and standards used were as recommended by the manufacturer. The data was analysed with MG-RAST using the default settings (Meyer et al., 2008).
3.3.6.8 Microbial Diversity Assessments of the Fissure Water using Visual Methods
Each time fissure water was collected, different staining and microscopy techniques were conducted in order to visualize the microorganisms in the water and to determine whether the cells were alive or dead.
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3.3.6.8.1 Negative Stain
Negative staining involved the addition of a small drop of the Nigrosin dye (1% w/v) to a clean, dry microscope slide. A drop of the fissure water was added to the Nigrosin, spread over the entire slide using a second microscope slide, and allowed to air dry. The stained cells were viewed with an Axioskop light microscope (Zeiss) under the 100x oil immersion magnification and images were captured using a DS-Fi1 (Nikon) camera with NIS Elements imaging software (V.3.22.00).
This stain can be used to indicate if a microorganism contains distinctive structures such as endospores, a flagellum, or a capsule (Clark, 1981; Green, 1990). The Nigrosin dye is negatively charged and therefore repelled by the negative charge of the microorganisms. This prevents the stain from entering the cell and staining the cytoplasm and as a result, the shapes and sizes of the microorganisms are seen as colourless shapes against the stained background. This stain can also be used to assess viability since the living cells will exclude the dye but it will enter the dead cells (Sanderson, 1994). Another advantage of using this stain is that fixation by heat or alcohol is not needed and therefore the microorganisms are seen in more life-like shapes.
3.3.6.8.2 Live/Dead Stain
A live/dead stain was conducted using the LIVE/DEAD® BacLightTM Bacterial Viability Kit (Molecular Probes, Inc., Invitrogen, Promega) according to the manufacturer’s instructions. The stained cells were viewed with an Axioskop light microscope (Zeiss) under the 100x oil immersion magnification and images were captured using a DS-Fi1 (Nikon) camera with NIS Elements imaging software (V.3.22.00).
This kit uses SYTO® 9 as the green-fluorescent nucleic acid stain and propidium iodide as the red-fluorescent nucleic acid stain. The SYTO® 9 stain enters all bacteria in the sample – the microorganisms with intact cell membranes and those with damaged cell membranes, whereas the propidium iodide enters only into the microorganisms with damaged cell membranes, causing a reduction in the SYTO® 9 stain fluorescence when both dyes are present. Therefore, microorganisms that are alive with intact cell membranes will stain green, while microorganisms that are dead or have damaged cell
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membranes will stain red when viewed under fluorescence. The excitation/emission for the dyes are 480/500 nm for the SYTO® 9 stain and 490/635 nm for the propidium iodide (Molecular Probes, Inc.).
3.3.6.8.3 Gram Stain
For the Gram-staining properties of the microorganisms, a Gram stain (Bartholomew & Mittwer, 1952) was conducted by heat fixing a drop of the fissure water onto a microscope slide. This was followed by flooding the entire slide with crystal violet for one minute and rinsing with dH2O. An iodine solution was then used to flood the slide for one minute and rinsed with dH2O. The decolourizer, ethanol (95% v/v) was added drop wise onto the slide, until the violet colour was no longer present and the slide was rinsed with dH2O. The counter stain, safranin red, was used by flooding the slide for one minute and rinsed with dH2O to remove any excess dye. Slides were air dried and the stained cells were viewed with an Axioskop light microscope (Zeiss) under the 100x oil immersion magnification and images were captured using a DS-Fi1 (Nikon) camera with NIS Elements imaging software (V.3.22.00).
The Gram stain differentiates microorganisms based on the chemical and physical properties of their cell walls through the detection of peptidoglycan – the basic component of the bacterial cell wall (Bartholomew & Mittwer, 1952).
3.3.6.8.4 Scanning Electron Microscopy
Scanning electron microscopy (SEM) and energy dispersive X-ray (EDS) analyses were conducted on the biome that was concentrated using TFF, as well as on the biomass that was attached to the sandstone in the PVC cartridges. For the fissure water samples, 100 ml of the water was filtered by collecting the biomass on a 0.2 µM Isopore Polycarbonate membrane filter (Merck Millipore). For the sandstone samples, a small piece of the crushed rock with attached biofilm was used.
The samples were fixed in 0.1 M sodium phosphate-buffered glutaraldehyde (3%, pH 7.0) for at least 3 hours, followed by a 1 hour fixation in similarly buffered osmium tetroxide (1%). The samples were dehydrated in a graded ethanol series (50%, 70% and 95%) for
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20 minutes in each phase. This was followed by two changes of 1 hour each, in 100% ethanol. The samples were dried using a critical point dryer (Tousimis), mounted onto 10 mm Cambridge aluminium stubs (Ted Pella Inc.) with epoxy glue and gold coated (60 nm) with a sputter coater (Bio-Rad). The specimens were examined with a JSM-7800F Extreme-resolution Analytical Field Emission SEM (FE-SEM) equipped with a 80 mm² X- Max energy dispersive X-ray analyses system (Oxford Instruments) and analysed using Aztec Software. SEM and EDS analyses were done at the Centre for Microscopy at the UFS (Bloemfontein, RSA).
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3.3.7 Summary of all Samplings and Collaborators
Table 3.4: A summary of the sampling conducted at the Star Diamonds mine.
Sampling: 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 Day: 21 24 24 06 29 18 19 20 31 28 23 06 11 26 28 16 15 20 Month: 03 03 04 06 06 07 07 07 08 01 04 06 03 06 07 10 12 01 Year: 2012 2012 2012 2012 2012 2012 2012 2012 2012 2013 2013 2013 2014 2014 2014 2014 2014 2015 On-site Physicochemical Analyses: pH Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N Temperature Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N Salinity Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N EC Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N ORP Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N DO Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N H2S Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N 3- PO4 Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N Fe2+ Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N Total Fe Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N H2O2 Y Y N Y Y Y Y Y Y Y Y Y Y Y N N N N Water Flow Rate Y Y N Y Y Y Y Y Y Y Y Y Y Y N Y Y N Gas Flow Rate N Y N Y N N N N N N N Y N N N N N N Water Chemistry Analyses: Anion/Cation Y N N N N N N N N N Y Y Y Y N N N N Formate/Acetate Y N N N N N N N N N N N N N N N N N Sulphur Species/Phosphorous Y N N N N N N N N N N N N N N N N N + NH3/NH4 Y N N N N N N N N N N N N N N N N N TN/DOC/DIC Y N N N N N N N N N N N N N N N N N Water Isotope Analyses: 14 δ CDIC Y N N N N N N N N N N N N Y N N N N δ18O and δD/δ2H Y N N N N N N N N N N N N N N N N N Gas Chemistry and Isotope Analyses: Serum Vials with Gas/Water N Y N N N N N N N N N Y N N N N N N Copper Tubes N Y N N N N N N Y N N Y N N N N N N Diversity: Cornelius® Canister Y N N Y Y Y Y Y Y Y Y Y Y Y Y N N N SterivexTM Filter N N N N I&R N N N N N N I&R N N N N N N Tangential Flow Filtration Y N Y N N N N N N Y Y N N N N N N N Massive Filter I R N N N N N N N N N N N I R N N N PVC Cartridges N N N N N N N N N N N N N N N N I R Legend: Y = Sample was taken, N = Sample was not taken, I = Filter was installed, R = Filter was retrieved.
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Table 3.5: The collaborators and institutions responsible for the analyses of selected samples.
Water Chemistry Analyses:
Anion Institute for Ground Water Studies, University of the Free State, Bloemfontein, RSA Cation Institute for Ground Water Studies, University of the Free State, Bloemfontein, RSA Formate/Acetate Princeton University, Princeton, New Jersey, USA Sulphur Species Princeton University, Princeton, New Jersey, USA Phosphorous Princeton University, Princeton, New Jersey, USA + NH3/NH4 Princeton University, Princeton, New Jersey, USA TN/DOC/DIC Princeton University, Princeton, New Jersey, USA / University of Toronto, Ontario, Canada Water Isotope Analyses:
14 δ CDIC Rafter Radiocarbon, National Isotope Centre, GNS Science, NZ δ18O and δD/δ2H University of Toronto, Ontario, Canada Gas Chemistry and Isotope Analyses:
Serum Vials with Gas/Water Princeton University, Princeton, New Jersey, USA / University of Toronto, Ontario, Canada Copper Tubes University of Bern, Bern, Switzerland
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3.4 RESULTS AND DISCUSSIONS
3.4.1 Fissure Water and Gas Analyses
3.4.1.1 On-Site Physicochemical Analyses
On-site analyses were conducted during sampling as discussed in Section 3.3.4.1, to monitor the basic geochemistry of the fissure water. These analyses were done over a period of 27 months, at random samplings, and are shown in Table 3.6.
Table 3.6: The basic on-site analyses of the fissure water measured over time.
Sampling: 1 2 4 5 6 7 8 9 Day: 21 24 06 29 18 19 20 31 Month: 03 03 06 06 07 07 07 08 Year: 2012 2012 2012 2012 2012 2012 2012 2012 Analyses: pH 8.45 8.65 8.27 8.21 8.18 8.41 8.41 8.7 Temperature (°C) 31.2 31.3 31.1 30.7 31.0 30.9 30.9 30.4 Salinity (g.L-1) 3 3 3 3 2 3 3 3 EC (mS.cm-1) 3.84 3.84 4.30 3.82 3.86 3.96 4.01 4.60 ORP (mV) -112 -131 -146 -226 -221 -210 -215 -168 DO (mg.L-1) 0.2 0.1 0.05 0.8 0.4 0.1 0.1 0.05 -1 H2S (mg.L ) 0 0.1 0.1 0.15 0.1 0.2 0.2 0.4 Fe2+ (mg.L-1) 0.1 0 <0.1 0 0 <0.1 0.1 <0.1 Total Fe (mg.L-1) <0.1 0 <0.1 0.4 0.1 <0.1 0.1 0.1 -1 H2O2 (mg.L ) 0 0 0 0 0 0 0 0 Water Flow Rate (L.min-1) 7.50 7.50 7.14 4.88 6.67 6.67 6.67 6.21 Gas Flow Rate (ml.min-1) - 207 167 - - - - - Sampling: 10 11 12 13 14 16 17 Day: 28 23 06 11 26 16 15 Month: 01 04 06 03 06 10 12 Year: 2013 2013 2013 2014 2014 2014 2014 Analyses: pH 8.66 8.4 8.48 8.33 8.32 - - Temperature (°C) 28.3 30.5 30.3 30.3 30.5 - - Salinity (g.L-1) 2.00 2.00 2.29 2.21 2.26 - - EC (mS.cm-1) 4.63 4.17 4.63 4.63 4.51 - - ORP (mV) -127 -94 -77 -104 -184 - - DO (mg.L-1) 0.05 0.1 0.1 0.8 0.8 - - -1 H2S (mg.L ) 0.4 0.4 0.4 0.2 0.5 - - Fe2+ (mg.L-1) <0.1 <0.1 <0.1 <0.1 <0.1 - - Total Fe (mg.L-1) <0.1 <0.1 0.2 <0.1 <0.1 - - -1 H2O2 (mg.L ) 0 0 0 0 0 - - Water Flow Rate (L.min-1) 4.39 5.46 4.5 3.33 2.73 3.33 3.53 Gas Flow Rate (ml.min-1) ------Legend: EC = electrical conductivity, ORP = oxidation reduction potential, DO = dissolved oxygen, “-“ = Not measured.
Throughout the time during which the borehole was monitored, the fissure water maintained a consistent moderate temperature and an alkaline pH, with an average of 30.6°C and 8.4, respectively. This is fairly similar for results obtained by Lin et al., (2006a)
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and Magnabosco et al., (2014), for the shallower fracture water samples. Although differences were observed in the ORP over time, the fissure water had a sustained reducing environment, with an average of -155 mV, even though the presence of dissolved
-1 -1 oxygen (0.3 mg.L ) was observed. The detection of H2S (0.24 mg.L ) was expected, due to the reducing environment. The EC was measured to estimate the total dissolved solids (TDS) in the fissure water, indicating a calculated average of 2.7 g.L-1, with moderate salinity of 2.6 g.L-1. Very low concentrations of Fe2+ (0.08 mg.L-1) and total Fe (0.12 mg.L-1) were observed and no H2O2 was detected, indicating that radiolysis of water could not be observed. The borehole had an average water flow rate of 5.6 L.min-1 and gas flow rate of 187 ml.min-1.
3.4.1.2 Geochemical Analyses
Geochemical analyses of the fissure water were conducted as discussed in Section 3.3.4.2, to also provide insight into the possible electron donor and acceptor conditions for biological cycling, and are shown in Table 3.7 and Figure 3.17. Determinands with values always below the detection limits are not shown.
Table 3.7: Geochemical analyses of the fissure water.
Sampling: 1 11 12 13 14 Day: 21 23 06 11 26 Month: 03 04 06 03 06 Year: 2012 2013 2013 2014 2014 Analyses: pH 8.46 8.16 7.71 7.35 7.87 EC (mS.m-1) 430 432 424 431 434 DOC (mg.L-1) 0.25 - - 0.69 0.64 DIC (mg.L-1) 5.3 - - - - TN (mg.L-1) 0.415 - - - - TOC (mg.L-1) - 1.00 1.07 0.97 1.53 TDS (mg.L-1) 2062 2301 2497 2223 2440 Lactate (mg.L-1) 0.025 - - - - HS- (mg.L-1) 2.87 ------1 NO3 (mg.L ) 0.049 - - - - + -1 NH4 (mg.L 0.27 - - - - P-Alkalinity (mg.L-1) 0 0 0 0 0 M-Alkalinity (mg.L-1 25 24.1 24.3 22 26.2 Calcium Hardness (mg.L-1) - 141.8 157.6 10 - Magnesium Hardness (mg.L-1) - 7.1 7.1 783 - Total Hardness (mg.L-1) - 148.9 164.7 793 - Aluminium as Al (mg.L-1) 0.18 0.014 0.022 0.037 0.024 Ammonia as N (mg.L-1) - - 0.562 0.84 0.617 Barium as Ba (mg.L-1) 0.035 - - - 0.718 Boron as B (mg.L-1) - - - 2.87 - Bromide as Br (mg.L-1) 5.06 6.8 6.54 6.47 9.11 Cadmium as Cd (mg.L-1) - - - - <0.001 Calcium as Ca (mg.L-1) 43 56.7 63.05 3.9 64.7 Chloride as Cl (mg.L-1) 963 1326 1467 1405 1497 Cobalt as Co (mg.L-1) 0.0013 - - - -
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Copper as Cu (mg.L-1) Results remained relatively constant over time and were fairly similar to data obtained from the shallower samples as discussed in Lin et al., (2006a). The fissure water had a high mineral content, was moderately saline and contained low sulphate, ammonium, and nitrate concentrations, having average values of 1332 mg.L-1 chloride, ~2.92 mg.L-1 sulphate, 0.27 mg.L-1 ammonium, and ~0.049 mg.L-1 nitrate. The DIC (5.3 mg.L-1) was significantly higher than the DOC (0.25 mg.L-1). The high DIC concentrations suggest high bicarbonate and CO2 levels in the fissure water, whereas the low DOC concentrations suggest low organic matter content. Very low concentrations of the trace elements were present in the fissure water. When the parameters were compared with a Durov diagram, created with The Geochemist’s Workbench® (V.10 Professional) software (Figure 3.17), it can be seen that the fissure water stayed geochemically very similar over time, indicating that influx from other water sources due to mixing was negligible for this period. Figure 3.17: A Durov diagram for the geochemical analyses of the fissure water. Chapter 3 75 3.4.1.3 Gas Chemistry and Isotope Analyses Analyses of the dissolved gas compositions in the fissure water were conducted as discussed in Section 3.3.4.3. Results obtained showed high concentrations of CH4 (82.7%), with trace amounts of O2 (0.01%), H2 (0.0004%), CO (0.0035%), CO2 (0.01%), and He (0.82%). The amount of N2 obtained was 15.79% and 1.44% for Ar. As for the C2H6, C3H8, iso- & n-C4H10, and iso- & n-C5H12 samples, the amounts obtained were all <0.01%. Onstott et al., (2006) indicated that the H2 is sustaining the subsurface microbial communities such as the H2-utilizing sulphate-reducing bacteria (SRB) and methanogens, and is produced by water radiolysis at a rate of ~1 XRS Myr−1. Therefore the importance of the detection of H2 in the fissure water is that it appears to be the most important energy source available to deep subsurface communities (Lin et al., 2005; Nealson et al., 2005). Isotope analyses of the gas and the fissure water were tested as discussed in Section 3.3.4.3 and results were added to the graphs from Ward et al., (2004) (Figure 3.18). The 2 13 δ HCH4 and δ CCH4 analyses yielded values of -190‰ and -51.3‰, respectively (Figure 14 3.18A). The δ CDIC yielded a value of -952‰ (~0.8), resulting in the fissure water having a calculated conventional radiocarbon age of 24 320 (~131) years, before present (BP) and the δ18O and δD/δ2H analyses yielded values of -6.27‰ and -38.64‰, respectively, falling on the global meteoric water line (GMWL) (Figure 3.18B), as described in Craig, (1961). According to Onstott et al., (2006), paleo-meteoric water ranges in age from ~10 Ka to >1.5 Ma, is of low salinity, falls along the GMWL and is rich in CO2 and atmospheric noble gas, therefore, the fissure water from the Star Diamonds mine represents a subsurface residence time consistent with the isotopic signatures that indicate fissure water of paleo- meteoric origin. These results are consistent with mixing between thermogenic and microbial gases, with positional and isotopic characteristics that are consistent with production by methanogens utilizing the CO2 reduction pathway (Sherwood Lollar et al., 2008), and indicates methane with a biogenic origin. All these isotopic analyses corresponded with results obtained by Ward et al., (2004); Onstott et al., (2006); Sherwood Lollar et al., (2008); Lau et al., (2014); and Magnabosco et al., (2014), for the samples collected at approximately the same depth as the borehole used in this research (Section 3.3.3). Chapter 3 76 A B 13 2 Figure 3.18: (A) δ CCH4 and δ HCH4 values for all samples compared to empirically determined fields for thermogenic gas and microbial gas produced by CO2 reduction (CR) and acetate fermentation (AF) as described in Whiticar, (1999) 2 18 and (B) δ HH2O and δ OH2O for fissure water samples. The global meteoric waterline (GMWL) is from Craig, (1961). In both graphs, samples from the Karoo formation (Middelbult mine) are indicated by (+). All samples from the Witwatersrand Supergroup are open symbols, and all samples from the Ventersdorp Supergroup are solid symbols. Symbols from individual mines are as follows: Merriespruit and Masimong (circles), Beatrix (squares), Evander (triangles), Kloof (diamonds). Kidd Creek data (X) are for abiogenic methane (Sherwood Lollar et al., 2002). (Obtained from Ward et al., 2004). Samples from the Star Diamonds mine are indicated by a red star. 3.4.2 Sandstone X-Ray Fluorescence Analyses Major-element XRF analyses were conducted as discussed in Section 3.3.5.2 on the sandstone from the Star Diamonds mine, before and after sterilization thereof. Results of the analyses are shown in in Table 3.8. Table 3.8: Major-element XRF analyses of the sandstone, before and after sterilization. Sandstone Sandstone Sandstone Sandstone Autoclaved & Acid Autoclaved Acid Washed Washed Analyses: Al2O3 (%) 20.81 20.131 20.123 20.142 Fe2O3 (%) 1.065 1.247 1.198 1.28 CaO (%) 0.064 0.051 0.031 0.035 MgO (%) 0.17 0.197 0.202 0.261 MnO (%) 0.012 0.06 0.055 0.061 Na2O (%) 0 0 0 0 K2O (%) 2.64 2.518 2.5 2.512 P2O5 (%) 0.039 0.045 0.045 0.043 TiO2 (%) 1.103 1.131 1.119 1.125 SiO2 (%) 74.01 74.304 74.363 74.316 LOI (%) 0.087 0.316 0.364 0.225 Total (%) 100 100 100 100 Chapter 3 77 All sandstone consists of the same general minerals that form the basis of the sandstones. These minerals are generally quartz and feldspars and consist mainly of silicon (Jackson, 1997; Boggs, 2000). From the XRF results obtained, it can be seen that the major components of the sandstone are silicon (74%) and aluminium (20%). The rest of the components are in the minority. Thus, the sterilization through autoclaving and acid washing did not appear to influence the percentages of the minerals that were tested for during the experiments. 3.4.3 Characterization and Diversity of the Deep Subsurface Microbial Biome 3.4.3.1 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis Although microbial diversity assessments were carried out on all the canisters, the SterivexTM filters and the TFF concentrated fissure water that were sampled over time (Table 3.4), only one agarose gel of each domain of life for the PCR, the nested DGGE PCR, and the DGGE are shown in the results as a representative of all the samples. 3.4.3.1.1 Genomic DNA Isolation and Polymerase Chain Reaction Genomic DNA was isolated from all the filters of the Cornelius® canisters, the SterivexTM filters, as well as the TFF concentrated fissure water as discussed in Section 3.3.6.6.1. Amplification of the Archaeal 16S (~600 bp) (Figure 3.19B), the Bacterial 16S (~1500 bp) (Figure 3.19C), and the Eukaryal 18S (~1700 bp) (Figure 3.19D) rRNA gene fragments were done as discussed in Section 3.3.6.6.2. Amplicons, of expected size for each primer set, were obtained for the rRNA gene fragments from all the samples. The bands of interest were excised from the agarose gel and purified (Section 3.3.6.6.2). Chapter 3 78 1 2 3 4 5 6 7 8 9 10 11 12 A B 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 C D Figure 3.19: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific), used to determine the size of the amplicons, with the amplified rRNA gene fragments for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The samples are shown in lanes 4 – 12. 3.4.3.1.2 Nested Polymerase Chain Reaction The purified PCR products were used to amplify the V3/V4 hypervariable regions for Archaea (~600 bp) (Figure 3.20B), Bacteria (~600 bp) (Figure 3.20C), and Eukarya (~600 bp) (Figure 3.20D) using a nested DGGE PCR as discussed in Section 3.3.6.6.3. Amplicons were obtained for the V3/V4 hypervariable regions from all the samples. Chapter 3 79 1 2 3 4 5 6 7 8 9 10 11 12 13 A B 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 C D Figure 3.20: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific), used to determine the size of the amplicons, with the amplified V3/V4 hypervariable regions for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The samples are shown in lanes 4 – 12. 3.4.3.1.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses The DGGE PCR products were used to construct a preliminary diversity profile for Archaea (Figure 3.21A), Bacteria (Figure 3.21B), and Eukarya (Figure 3.21C), through the use of DGGE (Section 3.3.6.6.4). Chapter 3 80 A B C Figure 3.21: DGGE diversity profiles for (A) Archaea, (B) Bacteria, and (C) Eukarya. From the DGGE assessments, it can be seen that all three domains of life (Archaea, Bacteria, and Eukarya) were present in the subsurface biome. This corresponded with results obtained by numerous researchers for various subsurface samplings at different locations, with some of them as described in Phelps et al., (1994); Takai et al., (2001); Moser et al., (2003); Kieft et al., (2005); Gihring et al., (2006); Lin et al., (2006a); Lin et al., (2006b); Onstott et al., (2009); Borgonie et al., (2011); Ragon et al., (2013); Lau et al., (2014); Magnabosco et al., (2014); and Mu et al., (2014). The DGGE results indicate that the Archaea and Eukarya profiles were consistent between the various sampling events, but differences were obtained within the Bacterial profiles. However, there are some drawbacks in using the DGGE technique, with one of Chapter 3 81 them being that only small fragments are separated through electrophoresis and may therefore limit the information obtained through sequencing and as a result, minor populations can be below the detection limit (Duarte et al., 2012). Another drawback is that different DNA sequences may have similar mobilities due to their identical GC content. Therefore, one band on a gel can represent more than one species. Also possible intra- specific or intra-isolate heterogeneity of the rRNA genes, as well as multiple copy numbers of the 16S rRNA operons of microorganisms can result in one species representing more than one band on a gel, giving an underestimation of the microbial diversity (Nakatsu et al., 2000; Muyzer et al., 2004; Michaelsen et al., 2006; Duarte et al., 2012). As a result, final conclusions about diversity cannot be made by just evaluating a DGGE gel alone, since results obtained can be misleading, but this technique can be used as a quick indication to show if there are any possible variations in the microbial community present in the sample, if it is kept in mind that there are shortcomings. Therefore, sequencing and analyses were done on selected DGGE bands as discussed in Sections 3.3.6.6.5 and 3.3.6.6.6. The analysed sequences indicated that the Bacteria were dominated by the Proteobacteria phylum, mainly consisting of the Alpha-, Beta-, and Gammaproteobacteria classes. This was followed by the Firmicutes phylum, consisting of the Bacilli and Clostridia classes and the Actinobacteria class of the Bacteriodetes phylum. For the Eukarya, the main phyla present were the Ascomycota, the Annelida, and the Basidiomycota. The Archaea were dominated by the Euryarchaeota phylum, which included the Methanococci, the Thermococci, the Methanobacteria, and the Methanomicrobia classes, followed by the Crenarchaeota containing the Thermoprotei class. These results corresponded with subsurface diversity results obtained by numerous researchers (Banfield et al., 1999; Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Moser et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013; Lau et al., 2014; Magnabosco et al., 2014). 3.4.3.2 Microbial Diversity Assessments using Targeted 16S and 18S rRNA Gene Sequencing The microbial diversity results discussed in this Section, are only a representation of the diversity that has been found using the different next generation sequencing techniques Chapter 3 82 and even though the rarefaction curve, indicating the annotated species richness (Figure 3.22E), did not reach a plateau, the reduced slope of the curve towards the end suggested that the sequences are representative of the microbial community. These results confirmed what has been shown with the DGGE analyses that microorganisms from all three domains of life (Archaea, Bacteria, and Eukarya) were found to be present within the biome of the fissure water (Figure 3.22). The biome was dominated by the Bacteria (>90%), but also contained a small fraction of Eukarya and Archaea. Various studies on subsurface biomes have concluded that the subsurface is generally dominated by the Bacteria and the microbial activities that dominate these extreme environments are fermentation, methanogenesis, and sulphate-reduction (Banfield et al., 1999; Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013). Extending the DGGE results, the Bacterial domain (Figure 3.23), was found to be dominated by the Proteobacteria phylum, followed by the Firmicutes and the Bacteroidetes. The Proteobacteria mainly consisted of Beta- and Gammaproteobacteria classes, followed by smaller fractions of the Delta- and Alphaproteobacteria. The majority of the Firmicutes were from the Bacilli and Clostridia classes. This is similar to what has been found by Moser et al., 2003; Lau et al., 2014; and Magnabosco et al., 2014; that the Proteobacteria is the most abundant phylum in the subsurface. For the Eukaryal domain (Figure 3.24), the main phyla present were Arthropoda, Streptophyta, Nematoda, Apicomplexa, and Chlorophyta. The majority of the Arthropoda consisted of the Insecta class and most of the Nematoda were of the Chromadorea class. The Archaeal domain (Figure 3.25) was dominated by the Euryarchaeota phylum, which include the methanogens and halobacteria, followed by the Thermoprotei which is a class of the Crenarchaeota. Other than the methanogens which can fix CO2, several sulphate-reducing bacteria were found amongst the Deltaproteobacteria such as the Desulfobacterales, Desulfovibrionales, and Syntrophobacterales orders (Muyzer & Stams, 2008). Amongst the microorganisms known to fix nitrogen, the Cyanobacteria and Chlorobi phyla, as well as the Pseudomonadales, Rhizobiales, and Actinomycetales orders were found to be present. This confirmed that microorganisms with various metabolisms were present in the subsurface biome (Figure 3.22). Chapter 3 83 A B (derived from Bacteria) (derived from Eukaryota) (derived from unclassified sequences) C E D ales acteria) (derived from Bacteria) uences) (derived from Bacteriodetes) Figure 3.22: MG-RAST taxonomic hit distribution of the diversity in the subsurface biome for (A) domain, (B) phylum, (C) Chapter 3 class, and (D) order, with (E) a rarefaction curve indicating the annotated species richness. 84 Figure 3.23: MG-RAST Krona diagram for the Bacteria at order level. Chapter 3 85 Figure 3.24: MG-RAST Krona diagram for the Eukarya at order level. Chapter 3 86 Figure 3.25: MG-RAST Krona diagram for the Archaea at order level. Chapter 3 87 3.4.3.3 Microbial Diversity Assessments of the Fissure Water using Visual Methods Microscopy images shown throughout this document are a representation of some of the diversity that has been found. Images were only used to indicate the various morphologies present, whether the microorganisms were alive or dead and what their Gram-staining properties are. These images should not be used as a quantification method for the amount of biomass obtained or to determine the ration of live and dead cells. The camera used to capture the images had a defect on its lens which is reflected on all the images. The defect can be seen in the bottom left corner of each image and should not be seen as a biological structure. 3.4.3.3.1 Staining The negative stain images (Figure 3.26) and the live/dead stain images (Figure 3.27) indicated that several diverse microorganisms with varying morphology and distinctive structures were present in the subsurface biome. It can also be seen that several macrostructures, as well as biofilm-like structures through the accumulation of cells were found and even though these structures were not identified, they are indicative of the complex nature of the biofilms found in the subsurface (Borgonie et al., 2011; Blanco et al., 2014). Biofilms are described as complex aggregations of microorganisms, strongly attached to a surface and encased within a self-produced extracellular matrix of polymeric substances (EPS). Biofilms are thought to be the major form of microbial life in natural environments, due to the biofilm offering structural support and protection from physical, chemical and biological stresses. Biofilms can also provide the optimal location relative to substrates required for metabolic function, as well as symbiotic benefits in multi- species communities (Watnick & Kolter, 2000; Costerton & Stewart, 2001; Stoodley et al., 2002; Mitchell et al., 2009; Lewandowski & Beyenal, 2013). Biofilms are characterized by structural heterogeneity, genetic diversity, and complex community interactions (Mitchell et al., 2009), and were likely formed by the microorganisms in the Chapter 3 88 subsurface to protect themselves from stresses created by the extreme conditions which the biomes encounter in the subsurface. Chapter 3 89 Figure 3.26: Negative stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm. Chapter 3 90 Figure 3.27: Live/dead stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm. Subsurface biomes in general are dominated by the Gram-negative Proteobacteria and the Firmicutes, which are mainly Gram-positive (Moser et al., 2003; Lin et al., 2006a; Lau et al., 2014; Magnabosco et al., 2014). This was confirmed for the subsurface biome from the Star Diamonds mine through the diversity results (Section 3.4.3.2), therefore, it was expected that microorganisms from both the Gram-positive and the Gram-negative groups should be present in the subsurface biome. This was confirmed by the Gram stain images (Figure 3.28). Chapter 3 91 Figure 3.28: Gram stain images of the fissure water from the Star Diamonds mine. All scale bars are equal to 2 µm. 3.4.3.3.2 Scanning Electron Microscopy Scanning electron microscopy (Figure 3.29) was conducted as discussed in Section 3.3.6.8.4. Images were falsely coloured using Adobe® Photoshop (V.CS5). EDS analyses confirmed that the biofilm consisted of biological material. Chapter 3 92 Figure 3.29: Scanning electron microscopy images of the fissure water and biofilm from the Star Diamonds mine. Scale bars are equal to either 10 µm or 1 µm. Chapter 3 93 3.5 CONCLUSIONS A study site was obtained in the Karoo sandstone within the Star Diamonds mine and comprehensive sampling was conducted of the fissure water to characterize the biogeochemistry of the study site over time. On-site analyses revealed that fissure water maintained a consistent moderate temperature and a slightly alkaline pH and even though differences were observed in the ORP over time, the fissure water had a sustained reducing environment. No H2O2 was detected, thus indicating no radiolysis in the water. Geochemical analyses of the fissure water were conducted to provide insight into the possible electron donor and acceptor conditions for biological cycling and also confirmed that the water chemistry remained fairly constant, possibly an indication that influx from older water sources are not relevant during this period. The fissure water had a high mineral content, was moderately saline and contained low sulphate, ammonium, and nitrate concentrations. The fissure water represented a subsurface residence time consistent with the isotopic signatures of paleo-meteoric origin and analyses of the conventional radiocarbon age for the fissure water was calculated at 24 320 (~131) years BP, with values falling on the global meteoric water line, again an indication that mixing with older water was negligible. The water also contained dissolved gases, including CH4, H2, and He. Their isotopic signatures were analysed and the results indicated a mixing of thermogenic and biogenic gases, with positional and isotopic characteristics that could be consistent with production by methanogens utilizing the CO2 reduction pathway. Over the past 20 years, comprehensive studies of subsurface diversity and geochemistry have revealed a wide diversity of microorganisms from all three domains of life in deep subsurface environments. These studies have shown that their metabolic activities can significantly contribute to the geochemistry of the deep subsurface, even though it happens orders of magnitude slower than for the microorganisms observed in surface environments (Phelps et al., 1994; Fredrickson & Onstott, 1996; Colwell et al., 1997; Onstott et al., 1998; Pedersen et al., 2000; Moser et al., 2003; Kieft et al., 2005; Gihring et al., 2006; Onstott et al., 2009; Borgonie et al., 2011; Ragon et al., 2013). The microbial diversity assessments of the fissure water from the Star Diamonds mine indicated that all three domains of life were present in the subsurface biome. The Chapter 3 94 Bacterial domain was found to be dominated by the Proteobacteria phylum, followed by the Firmicutes. The Proteobacteria mainly consisted of Alpha-, Beta-, and Gammaproteobacteria classes and the majority of the Firmicutes were from the Bacilli and Clostridia classes. The Archaeal domain was dominated by the Euryarchaeota phylum, which includes the methanogens and halobacteria, followed by the Thermoprotei, which is a class of the Crenarchaeota. For the Eukaryal domain, the main phyla present were Arthropoda, Streptophyta, Nematoda, Apicomplexa, and Chlorophyta. This confirmed that microorganisms which grow optimally under chemolithoautotrophic conditions, in the presence of H2 and CO2 (as many biomes, from different subsurface environments encounter), as well as microorganisms with heterotrophic metabolisms were present in the subsurface biome and included methanogens, which can fix CO2, sulphate-reducing bacteria, and microorganisms known to fix nitrogen. Several macrostructures, as well as biofilm-like structures, were found in the subsurface biome, and even though these structures were not identified, it is indicative of the complex nature of the biofilms found in the subsurface. 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Chapter 3 108 CHAPTER 4 MONITORING OF GROWTH AND LOW PRESSURE STUDIES 4.1 INTRODUCTION The diversity studies of the subsurface biome from the Star Diamonds mine (Chapter 3) displayed an extensive variety in the microbial community and included species from all three domains of life. However, limited, detailed reports of subsurface biome survival, community shifts, or metabolic cycling under low pressures are available (Morozova et al., 2010; Glossner, 2013), since previous research studies under pressure usually focussed on the characterization of single microorganisms (Miller et al., 1988; Nelson et al., 1992; Bothun et al., 2004; Boonyaratanakornkit et al., 2007; Takai et al., 2008; Hiraki et al., 2012). According to Fakruddin & Mannan (2013), two important questions to address when studying a microbial community are how do the microorganisms function and how does the composition of the community adjust due to stress conditions from environmental changes? These changes are usually harmful to the microorganisms and may lead to an overall decrease in metabolic activity, turnover times, and cell density (Phelps et al., 1994; Teske, 2005; Onstott et al., 2009). Therefore, knowledge regarding the survival of microorganisms within a subsurface biome that are subjected to environmental changes such as different physiological growth conditions with differences in nutrition and the addition of lower pressures, are required to enhance the current literature. As a result, studies of viable cultures that can maintain subsurface cycling are required to investigate how CCS conditions will introduce additional impacts on the biome. 4.1.1 Eubacterium limosum E. limosum is an obligate anaerobic, non-sporulating, acetogenic bacterium from the family, Eubacteriaceae (Song & Cho, 2015). It belongs to the Clostridiales order under the Clostridia class, the Firmicutes phylum, and the Bacteria kingdom. They are Gram- Chapter 4 109 positive rods that contain a thick peptidoglycan layer in their cell wall (Song & Cho, 2015), and are generally isolated from anaerobic digesters (Roh et al., 2011), rumen fluid of sheep (Genthner et al., 1981), sewage (Genthner et al., 1981), or human faeces (Finegold et al., 1983). Due to the anaerobic metabolism of E. limosum and the fact that it grows optimally under chemolithoautotrophic conditions in the presence of H2 and CO2 [as many biomes from different subsurface environments encounter (Lovley & Chapelle, 1995; Stevens, 1997; White et al., 1998; Lin et al., 2005; Onstott et al., 2006; Porter et al., 2009; Woycheese et al., 2015)], and even under two bar pressure (Genthner et al., 1981; Genthner & Bryant, 1982; Drake, 1994; Lelait & Grivet. 1996), it was used as the positive control organism for this research to benchmark growth under pressure. E. limosum is able to grow on a variety of substrates, including hydrogen (H2), carbon monoxide (CO), carbon dioxide (CO2), glucose, and methanol (Genthner et al., 1981; Loubière & Lindley, 1991; Lelait & Grivet, 1996; Roh et al., 2011). This microorganism has been considered a microbial syngas catalyst due to its high growth rate under pressures above 1 atm (1.01325 bar), when using CO as the sole carbon or energy source (Roh et al., 2011). Through the use of the Wood–Ljungdahl pathway [also known as the reductive acetyl coenzyme A (Acetyl-CoA) pathway], E. limosum can fix CO and CO2 to produce Acetyl-CoA which is then converted into acetate, butyrate, or ethanol as fermentation products and has therefore been considered a model strain for bio-energy production from syngas (Loubière & Lindley, 1991; Roh et al., 2011). 4.2 AIMS OF THIS CHAPTER The main aims of this chapter were to: Study the potential growth of the subsurface biome from the Star Diamonds mine under different physiological growth environments while assessing diversity changes. Introduce lower pressure to the microorganisms in order to determine if they can survive these conditions. Chapter 4 110 4.3 MATERIALS AND METHODS Unless otherwise specified, the anaerobic techniques used throughout this research were according to the procedures of Bryant, (1972); Hungate & Macy, (1973); Balch & Wolfe, (1976); and Boone & Bryant, (1980); and all incubations for cultivation purposes of the subsurface biome and E. limosum were conducted at 35°C in a temperature controlled incubator. Even though this temperature was slightly higher than the fissure water (Section 3.4.1.1), it was to accommodate the optimum growth temperature for E. limosum. This was also selected to ensure that the CO2 would be in a supercritical state during CCS conditions. All growth curves were constructed using GraphPad Prism® (V.5.03) (GraphPad Software, Inc.), by plotting the mean and error values from three replicates of independent growth studies on a logarithmic scale. The rate of growth per unit amount of biomass [also known as the maximum specific growth rate (µmax)], was calculated using (Equation 4.1) (Pirt, 1975). This was determined by linear regression analyses with Microsoft® Excel (Microsoft® Corporation) of the exponential phase of the growth curve. The period of time required for the population to double [also known as the doubling time (td)], was calculated using (Equation 4.2) (Pirt, 1975). The µmax and td was calculated for each individual growth curve, followed by obtaining the average of three replicates. The average values with standard deviations (SD) were given in the results. (Equation 4.1) (µmax) = maximum specific growth rate (x0) = initial biomass concentration (xt) = biomass concentration at the time given (t) = time in hours (Equation 4.2) td = ln(2)/µmax (td) = doubling time (µmax) = maximum specific growth rate Chapter 4 111 4.3.1 Medium Preparation for the Positive Control Microorganism Eubacterium limosum (DSM-20543) was obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ) culture collection as a lyophilized pellet. The culture was revived using the anaerobic PYG medium (medium 104) from the DSMZ (5 g.L-1 trypticase peptone, 5 g.L-1 peptone, 10 g.L-1 yeast extract, 5 g.L-1 beef extract, 5 -1 -1 -1 -1 -1 g.L glucose, 2 g.L K2HPO4, 1 ml.L Tween 80, 0.5 g.L cysteine-HCl.H2O, 1 mg.L -1 -1 -1 resazurin, 40 ml.L salt solution, 10 ml.L haemin solution, 0.20 ml.L vitamin K1 -1 solution, pH 7.2). The salt solution was prepared by dissolving 0.25 g.L CaCl2.2H2O, 0.5 -1 -1 -1 -1 -1 g.L MgSO4.7H2O, 1 g.L K2HPO4, 1 g.L KH2PO4, 10 g.L NaHCO3, and 2 g.L NaCl -1 in dH2O. For the haemin solution, 50 mg haemin was dissolved in 1 N NaOH (40 g.L ) and made up to 100 ml with dH2O. For the vitamin K1 solution, 0.1 ml of vitamin K1 was dissolved in 20 ml EtOH (95%) and filter-sterilized. The vitamin K1 solution, the haemin solution, and the cysteine-HCl.H2O were only added after the medium has been boiled for 5 minutes and cooled for 10 minutes, while flushing with CO2 to decrease the oxygen solubility. The pH was adjusted to 7.2 using 8 N NaOH (320 g.L-1) and the medium was dispensed into serum vials that were closed with blue chlorobutyl rubber stoppers (Bellco Glass Inc.) and sealed with aluminium crimp caps (Lasec). The medium was degassed for one hour using nitrogen (N2) gas (99.999%) and subsequently sterilized. The resazurin was added as an oxidation-reduction indicator and the cysteine-HCl.H2O was added to remove the remaining oxygen (Sambrook et al., 1989). To revive the culture, the ampoule containing the lyophilized pellet, was opened under sterile conditions inside a laminar flow cabinet (Labotec®) and the pellet was transferred in a sterile 1.5 ml eppendorf tube and added, under anaerobic conditions, to a 150 ml serum vial containing 80 ml of the PYG medium. This was done in an anaerobic glove box (Coy Laboratories) that was flushed three times at 200 kPa (2 bar) with nitrogen gas (99.999%), followed by two times with a combination of N2, CO2 and H2 (80:10:10 v/v/v). 4.3.2 Identification of the Positive Control Microorganism Molecular identification was conducted on E. limosum (DSM-20543). Chapter 4 112 4.3.2.1 Genomic DNA Isolation and Polymerase Chain Reaction Genomic DNA was isolated from E. limosum using the NucleoSpin® Soil kit (Macherey- Nagel) as discussed in Section 3.3.6.6.1. A PCR was conducted to amplify the Bacterial 16S rRNA gene (~1500 bp) (Section 3.3.6.6.2) using the universal oligonucleotide primers for Bacteria (Section 3.3.6.6.2, Table 3.1). The PCR products were visualized on a 1% (w/v) agarose gel and purified (Section 3.3.6.6.2). 4.3.2.2 Ligation of the 16S rRNA Gene The purified PCR product was ligated into the pGEM®-T Easy Vector system (Promega) (Figure 4.1), as per manufacturer’s instructions. The ligation reaction was performed in a total reaction volume of 10 µl and consisted of 2x Rapid Ligation Buffer (5 µl), pGEM®-T Easy Vector (2.5 ng), PCR product (3 µl), and T4 DNA Ligase (0.3 Weiss units). The ligation reaction was incubated overnight at 4°C. Figure 4.1: The pGEM®-T Easy Vector map and sequence reference points (Promega). Chapter 4 113 4.3.2.3 Transformation of the 16S rRNA Gene Competent Escherichia coli (E. coli) One Shot TOP10 (Invitrogen) cells were prepared according to the procedure of Hanahan, (1983), with modifications. Erlenmeyer flasks -1 -1 -1 containing 100 ml PSI broth (5 g.L yeast extract, 20 g.L tryptone, 5 g.L MgSO4.7H2O, pH 7.6 with KOH) were inoculated with 1 ml of an overnight culture and grown at 37°C on a rotary shaker (200 rpm), until an optical density (OD) of 0.6 at 600 nm was reached. This was then placed on ice for 30 minutes and centrifuged (3 000 x g, 10 minutes, 4°C), to collect cells. The cells were resuspended in 40 ml transformation buffer I (TfbI) (30 mM potassium acetate, 100 mM rubidium chloride, 10 mM CaCl2, 15% glycerol, pH 5.8) and incubated on ice for 15 minutes. Cells were collected through centrifugation (3 000 x g, 10 minutes, 4°C) and resuspended in 4 ml transformation buffer II (TfbII) (10 mM MOPS-NaOH, 75 mM CaCl2, 10 mM rubidium chloride, 15% glycerol, pH 6.5). The cell suspension was incubated on ice for 10 minutes and aliquots were made that were snap frozen in liquid nitrogen followed by storage at -80°C until used. Aliquots (50 µl) of the competent cells were removed and allowed to thaw on ice. The ligated plasmid reactions (5 µl) were added to the cells and incubated on ice for 30 minutes. The cell suspension was heat shocked for 40 seconds at 42°C and immediately placed back in ice-water for 2 minutes. SOC medium (250 µl) (20 g.L-1 tryptone, 5 g.L-1 yeast extract, 0.01 M NaCl, 0.0025 M KCl, 0.01 M MgCl2, 0.01 M MgSO4, 0.02 M glucose) was added to the transformation reactions and incubated for 60 minutes at 37°C, with gentle shaking. Different volumes were plated on separate Lysogeny broth (LB) plates (10 g.L-1 peptone, 5 g.L-1 yeast extract, 10 g.L-1 NaCl, 18 g.L-1 agar, pH 7.0), containing 100 µg.ml-1 ampicillin, 0.2 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and 40 µg.ml-1 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranosidehosphate (X-Gal) and incubated overnight at 37°C to verify DNA ligation using blue/white selection. Single white colonies were selected from the LB-AIX plates and separately inoculated into 5 ml LB medium containing 60 µg.ml-1 ampicillin and incubated overnight on a rotary shaker (200 rpm), at 37°C. Plasmids were isolated using the BioFlux BioSpin Plasmid DNA Extraction Kit (Bioer Technology Co., Ltd) according to the manufacturer’s instructions. Chapter 4 114 4.3.2.4 Analyses of the 16S rRNA Gene Double digestion was conducted on the plasmids using the restriction enzymes EcoRI and NdeI (Fermentas). Before the digestion was done, the 16S rRNA gene sequence of E. limosum ZL2, with accession number JQ085755.1 was retrieved from the Genbank Database (NCBI, 2012) and in silco analyses were conducted using the Restriction Mapper (V.3.0) tool, to confirm that the chosen restriction enzymes do not digest the 16S rRNA gene. The digestion reactions were performed in a total reaction volume of 50 µl and consisted of 10 µl purified plasmid, 5 µl buffer Tango (2x), 2.5 µl of each restriction enzyme, and sterile dH2O. The reaction mixture was incubated at 37°C for 3 hours, where after the enzymes were thermally deactivated at 65°C for 20 minutes. The restriction digestions were visualized on a 1% (w/v) agarose gel and purified (Section 3.3.6.6.2). Sequencing and analyses of the purified fragments were done as discussed in Section 3.3.6.6.6 using the primer set indicated in Table 4.1. Table 4.1: The RNA polymerase promoter primers used for sequencing. Primer Nucleotide sequence Tm (°C) T7 Promoter 5'- TAA TAC GAC TCA CTA TAG GG -3' 47 SP6 Promoter 5'- TAC GAT TTA GGT GAC ACT ATA G -3 42 4.3.3 Cultivation Conditions for Eubacterium limosum Since E. limosum is described in the literature to be an obligate anaerobe (Song & Cho, 2015), cultivation under aerobic conditions was not done. To study the growth of E. limosum under anaerobic conditions, the nutritionally rich, non-selective PYG medium was prepared as discussed in Section 4.3.1. An additional evaluation was conducted to test the ability of E. limosum to grow in the fissure water (FW), from the Star Diamonds mine. This was done with and without the addition of sandstone. The fissure water was used as a minimal medium, containing no additional nutrients and the sandstone was added as a potential nutrient source, since the rock and fissure water will be the Chapter 4 115 available nutrient source for the microorganisms in the subsurface. Both the PYG and -1 -1 FW contained 0.5 g.L cysteine-HCl.H2O and 1 mg.L resazurin and the PYG was prepared using fissure water. Hungate tubes (20 ml), containing 4 ml of either PYG or FW, were prepared, sealed with a blue chlorobutyl rubber stopper (Bellco Glass Inc.) and degassed for one hour using nitrogen gas (99.999%). The media were sterilized where after inoculums (10% v/v) were made with E. limosum under anaerobic conditions (Section 4.3.1). For the FW tubes that contained sandstone, 0.5 g crushed sandstone (Section 3.3.5) was added to the media before being sterilized. Negative controls were prepared for each inoculum. The tubes were incubated on a rotary shaker (200 rpm), at 35°C. For the time zero reading, a tube was immediately inserted into a photoLab S6 photometer [Wissenschaftlich-Technische Werkstätten (WTW), Labotec®] and the OD was measured at a set 605 nm. This was repeated at each time interval. Before the Hungate tubes were prepared, an OD605 of the tube alone was taken using the photometer and this value was subtracted. Growth studies continued for 48 hours and were done in triplicate for each media type. The OD values were plotted against the time intervals to obtain the various growth curves. Visual assessments were conducted on each completed growth study. In order to study the growth of E. limosum under pressure, low pressure were introduced. In the literature, when microorganisms are grown under pressure, several different gas compositions are used, some of them being H2:CO2 (80%:20% v/v) (Genthner et al., 1981; Girbal et al., 1997); H2:CO2 (60%:40% v/v) (Leclerc et al., 1997); N2:CO2 (90:10% v/v) (Men et al., 2014); N2:CO2 (85%:15% v/v) (Loubière & Lindley, 1991); N2:CO2 (80%:20% v/v) (Krumholz et al., 1996; Girbal et al., 1997; Bainotti et al., 1998); N2:CO2 (60%:40% v/v) (Genthner & Bryant, 1982) and N2:CO2:H2 (80%:10%:10% v/v/v) (Girbal et al., 1997). For the purpose of this research Section, the H2:CO2 combination was used. This also mimics conditions where the methanogens, and other anaerobic physiological types dominate the deep subsurface, survive (Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Onstott, 2005; Basso et al., 2009; Morozova et al., 2010). Chapter 4 116 Hungate tubes were again prepared as discussed previously for PYG and for the FW, with and without the addition of sandstone, for the inoculums, as well as for the negative controls. Each tube was pressurized to 2 bar (Figure 4.2A) with a gas ratio of 80% H2 and 20% CO2 (v/v) (Genthner et al., 1981), while using protective gear. Within the 2 bar, 20% (0.2 fraction of the total) had to be CO2. This meant that the tubes were pressurized, without releasing gas, to 0.4 bar using CO2 gas (99.99%) (2 bar x 0.2 fraction = 0.4 bar). To obtain this pressure, the CO2 regulator was set to 40 kPa (0.4 bar). The other 80% (0.8 fraction of the total) had to be H2, which meant that the tubes were further pressurized, without releasing gas, to 1.6 bar using H2 gas (99.999%) (2 bar x 0.8 fraction = 1.6 bar). To obtain this pressure, the H2 regulator was set to 200 kPa (2 bar). To pressurize the tubes, sterile syringe-tipped, high pressure Masterflex® Norprene hoses (Cole-Parmer®) were connected to the regulators (Figure 4.2B and C). A sterile 0.2 µM syringe filter and a sterile needle were connected to the syringe (Figure 4.2D). To insert the CO2, the gas tank was opened with the regulator set on 40 kPa (0.4 bar) and the hose was flushed for 30 seconds. The needle was inserted into the tube while the gas was flowing. The CO2 was allowed to enter into the tube for five seconds, where after the tube was inverted for six seconds to allow the gas to equilibrate between the hose and the tube. The presence of bubbles was an indication that gas was entering the media. The tube was turned back and the needle was removed with the gas still flowing. The hose was flushed again for ten seconds before the next tube was pressurized with CO2. To insert the H2, the gas tank was opened with the regulator set on 200 kPa (2 bar) and the hose was flushed for 30 seconds. The needle was inserted into the tube, while the gas was flowing. The H2 was allowed to enter into the tube for six seconds, where after the tube was inverted to ensure that gas was no longer entering and the needle was removed with the gas still flowing. The hose was flushed again before the next tube was pressurized with H2. Before a needle was inserted into the tube, the blue chlorobutyl rubber stopper was first sterilized, by flaming it with 70% ethanol (EtOH). The pressurized tubes were incubated on a rotary shaker (200 rpm), at 35°C and the OD605 was measured as discussed previously using a photoLab S6 photometer [Wissenschaftlich-Technische Werkstätten (WTW), Labotec®]. Growth studies continued for 48 hours and were done in triplicate for each media type. The OD values were plotted Chapter 4 117 against the time intervals to obtain the various growth curves and visual assessments were conducted on each completed growth study. C A B D Figure 4.2: (A) A Hungate tube pressurized to 2 bar and (B-D) the syringe-tipped, high pressure hoses used to pressurize the tubes. E. limosum cultures were stored at mid exponential phase (~18 hours), by preparing Hungate tubes (10 ml) with 2.5 ml glycerol (50% w/v), 7.5 µl resazurin (0.1% w/v), and 20 µl cysteine-HCl.H2O (2.5% w/v) to each tube. The glycerol was degassed for one hour using nitrogen (N2) gas (99.999%) and subsequently sterilized. The culture (2.5 ml) was added into each Hungate tube under anaerobic conditions (Section 4.3.1), and stored at -80°C. 4.3.4 Cultivation Conditions for the Subsurface Biome To study the growth of the subsurface biome from the Star Diamonds mine under aerobic conditions, the nutritionally rich, non-selective LB medium (10 g.L-1 peptone, 5 g.L-1 yeast extract, 10 g.L-1 NaCl, pH 7.0) (Sambrook et al., 1989) was prepared aerobically using fissure water. The additional evaluation of the ability of the subsurface biome to grow in the fissure water alone was again done, with and without the addition of sandstone. Erlenmeyer flasks (1 L) containing 180 ml of the LB medium were prepared Chapter 4 118 and sterilized. Inoculums (10% v/v) were made under aerobic conditions using the fissure water containing the TFF concentrated subsurface biome (Section 3.3.6.3). When the fissure water alone was used as the growth medium, 200 ml of the concentrated subsurface biome was used directly. For the FW flasks that contained sandstone, 20 g crushed sandstone (Section 3.3.5) was added to the media before sterilization. Negative controls were prepared for each inoculum. The flasks were incubated on a rotary shaker (200 rpm), at 35°C. For the time zero reading, 1 ml was immediately withdrawn and the OD was measured at 600 nm using a Spectronic® GenesysTM 5 spectrophotometer (Spectronic Instruments). This was repeated at each time interval. Growth studies continued for 32 hours and were done in triplicate for each media type. The OD values were plotted against the time intervals to obtain the various growth curves and microbial diversity assessments were conducted on each completed growth study using visual methods and molecular techniques. To study the growth of the subsurface biome from the Star Diamonds mine under anaerobic conditions, as well as under low pressures, LB medium and FW, with and without the addition of sandstone, were prepared anaerobically in 20 ml Hungate tubes as discussed in Section 4.3.3 and the inoculums (10% v/v) were made with the TFF concentrated subsurface biome (Section 3.3.6.3). When the fissure water alone was used as the growth medium, 5 ml of the concentrated subsurface biome was used directly. For the pressure studies, the tubes were pressurized to 2 bar using a gas ratio of 80% H2 and 20% CO2 (v/v) (Section 4.3.3). The tubes were incubated and measured at OD605 as discussed in Section 4.3.3. Growth studies continued for 48 hours and were done in triplicate for each media type. Microbial diversity assessments were again conducted on each completed growth study using visual methods and molecular techniques. 4.3.5 Microbial Diversity Assessments of Eubacterium limosum and the Subsurface Biome using Visual Methods Different staining and microscopy techniques were conducted on all the completed growth studies in order to assess whether any changes in diversity occurred, as well as Chapter 4 119 whether the cells were alive or dead. This included a negative stain using the Nigrosin dye as discussed in Section 3.3.6.8.1, a live/dead stain using the LIVE/DEAD® BacLightTM Bacterial Viability Kit (Molecular Probes, Inc., Invitrogen, Promega) as discussed in Section 3.3.6.8.2, and a Gram Stain as discussed in Section 3.3.6.8.3. 4.3.6 Microbial Diversity Assessments of the Subsurface Biome using Denaturing Gradient Gel Electrophoresis DGGE analyses were conducted on the completed growth studies with the subsurface biome to assess whether any changes in diversity occurred under the various conditions. 4.3.6.1 Genomic DNA Isolation and Polymerase Chain Reaction Genomic DNA was isolated using the NucleoSpin® Soil kit (Macherey-Nagel) as discussed in Section 3.3.6.6.1. A PCR was conducted to amplify the Archaeal 16S (~600 bp), Bacterial 16S (~1500 bp), and Eukaryal 18S (~1700 bp) rRNA gene fragments as discussed in Section 3.3.6.6.2 using the universal oligonucleotide primers for each gene (Section 3.3.6.6.2, Table 3.1). The PCR products were visualized on a 1% (w/v) agarose gel and purified (Section 3.3.6.6.2). 4.3.6.2 Nested Polymerase Chain Reaction A nested DGGE PCR was conducted on all the purified samples to amplify the V3/V4 hypervariable regions for Archaea (~600 bp), Bacteria (~600 bp), and Eukarya (~600 bp) as discussed in Section 3.3.6.6.3 using the DGGE oligonucleotide primers for each gene (Section 3.3.6.6.3, Table 3.2). The PCR products were visualized on a 1% (w/v) agarose gel. Chapter 4 120 4.3.6.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses The products from the nested DGGE PCR were used to construct preliminary diversity profiles using the DGGE DcodeTM universal mutation detection system (Bio-Rad Laboratories, Inc.) as discussed in Section 3.3.6.6.4. Selected bands were excised from the gel and the concentrations of the recovered DNA were increased through re- amplification of the V3/V4 hypervariable regions from the eluted DGGE products using the oligonucleotide primers as indicated in Section 3.3.6.6.5, Table 3.3. The re-amplified PCR products were visualized on a 1% (w/v) agarose gel and purified (Section 3.3.6.6.2). Sequencing and analyses of the purified fragments were done as discussed in Section 3.3.6.6.6. Chapter 4 121 4.4 RESULTS AND DISCUSSIONS 4.4.1 Molecular Identification of the Positive Control Microorganism 4.4.1.1 Genomic DNA Isolation, Polymerase Chain Reaction and Transformation of the 16S rRNA Gene The gDNA isolated from E. limosum (Section 3.3.6.6.1), was used to amplify the Bacterial 16S (~1500 bp) rRNA gene fragment (Figure 4.3) as discussed in Section 3.3.6.6.2. 1 2 3 4 5 A B Figure 4. 3: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, and (B) the amplified 16S rRNA gene fragment from E. limosum. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown and in lanes 2 – 5, the 16S rRNA gene fragment. Amplicons were obtained and the bands of interest were excised from the agarose gel and purified as discussed in Section 3.3.6.6.2. The purified PCR products were ligated into the pGEM®-T Easy Vector system (Promega) as discussed in Section 4.3.2.2, and transformed into E. coli One Shot TOP10 (Invitrogen) competent cells (Section 4.3.2.3). 4.4.1.2 Analyses of the 16S rRNA Gene The insert was excised with a double digestion reaction (Figure 4.4B) using the restriction enzymes EcoRI and NdeI (Fermentas) as discussed in Section 4.3.2.4. Chapter 4 122 1 2 3 4 5 A B Figure 4.4: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the inserts, and (B) the double digestion profile for E. limosum. In lane 1, the O’GeneRulerTM DNA Ladder Mix are shown and in lanes 2 – 5, the ~3000 bp plasmid backbone and the ~1500 bp insert. Positive ligation of the insert into the plasmid was obtained with the presence of the ~3000 bp plasmid backbone and the ~1500 bp insert. The inserts were purified (Section 3.3.6.6.2) and analyses were done as discussed in Section 4.3.2.4. Using the first five BLAST results, it was confirmed that the positive control microorganism were E. limosum (Table 4.2). Table 4.2: BLAST results for the 16S rRNA gene sequence of E. limosum. Accession Description Identity E-Value Eubacterium limosum gene for 16S ribosomal LC019782.1 99% 0.0 RNA, partial sequence, strain: JCM 9978 Eubacterium limosum strain ZL2 16S ribosomal JQ085755.1 99% 0.0 RNA, partial sequence Eubacterium limosum strain JCM 6421 16S NR_113248.1 99% 0.0 ribosomal RNA gene, partial sequence Eubacterium limosum strain Tr5 16S ribosomal HM590802.1 99% 0.0 RNA gene, partial sequence Eubacterium limosum KIST612 strain KIST612 NR_074533.1 99% 0.0 16S ribosomal RNA, complete sequence Chapter 4 123 4.4.2 Experimental Layout for the Cultivation of Eubacterium limosum and the Subsurface Biome Eubacterium limosum A A: B: Anaerobic Low Pressure 1: 2: 3: 1: 2: 3: PYG FW + Sandstone FW - Sandstone PYG FW + Sandstone FW - Sandstone Subsurface Biome B C&D: E: Aerobic & Anaerobic Low Pressure 1: 2: 3: 1: 2: 3: LB FW + Sandstone FW - Sandstone LB FW + Sandstone FW - Sandstone Figure 4.5: A summary of the experimental layout for the cultivation of (A) E. limosum and (B) the subsurface biome. Chapter 4 124 4.4.3 Cultivation and Assessments of Eubacterium limosum Growth curves were obtained for E. limosum by analysing the various growth studies under anaerobic conditions and low pressure for 48 hours (Figure 4.6) as discussed in Section 4.3.3. The µmax and td were calculated for growth in the PYG medium. 10 A1: PYG A2: FW + SS A3: FW - SS B1: PYG 605 1 B2: FW + SS OD B3: FW - SS 0.1 0 4 8 12 16 20 24 28 32 36 40 44 48 Time (Hours) Figure 4.6: Growth curves for E. limosum under anaerobic conditions (A) in blue and low pressure (B) in purple (FW = fissure water, +SS = with sandstone, -SS = without sandstone). From the results obtained, it can be seen that the pressure did not influence the growth of E. limosum, since growth curves were similar for the same type of medium used when anaerobic conditions were compared with low pressure. It can also be seen that exponential growth did not occur when the fissure water was used, indicating that the fissure water on its own did not provide all the necessary nutrients for E. limosum to grow optimally. The slight increase in the starting OD605 for the fissure water with sandstone (occurring for all the growth curves that contained sandstone throughout this chapter) was due to the weathering of the sandstone when in contact with the liquid. These sandstone particles are then detected when measuring the OD, but cannot be seen as growth and therefore it can be concluded that the presence of sandstone also had no effect on the growth, as the trends of the growth curves were similar for the fissure water, with and without the addition of sandstone. Chapter 4 125 As expected, the best biomass yields were obtained when E. limosum was grown in the -1 PYG medium. A µmax of 0.090 h and a td of 7.68 hours were calculated for growth under -1 anaerobic conditions, with a µmax of 0.071 h and a td of 9.81 hours for growth under low pressure. This indicated that growth under anaerobic conditions was slightly faster than growth under low pressure. In the literature, several different maximum growth rates are available for E. limosum, depending on which substrate was used for studying its growth under specific conditions (Mariotto et al., 1989; Le Bloas et al., 1993). However, the values for the maximum growth rates obtained in this study are similar to what Loubière & Lindley, (1991) described when studying the use of acetate as an additional co- substrate to improve the growth of E. limosum when CO2 fixation is rate-limiting. They found that when E. limosum was grown as a batch culture on methanol/CO2 in a medium supplemented with acetate, a short exponential growth phase was observed with a µmax of 0.12 h-1, followed by an extended period of decelerating growth. However, when E. limosum was grown in the absence of acetate (similar to growth in the PYG medium), -1 growth proceeded at a lower maximum growth rate, having a µmax of 0.075 h . Additional assessments were conducted on all the growth studies using different staining and microscopy techniques as discussed in Section 3.3.6.8. The negative controls remained sterile. From the negative stain images (Figure 4.7), it can be seen that biomass was obtained for all the parameters tested and that the cells were alive. A1: PYG B1: PYG Chapter 4 126 A2. FW + SS B2. FW + SS A3: FW - SS B3. FW - SS Figure 4.7: Negative stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm. The live/dead stain images (Figure 4.8) confirmed the previous observations and that most cells stayed alive under all the parameters tested. A1: PYG B1: PYG Chapter 4 127 A2: FW + SS B2. FW + SS A3: FW - SS B3. FW - SS Figure 4.8: Live/dead stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm. The Gram stain images (Figure 4.9) confirmed that E. limosum are, as described in the literature, a pure culture of Gram-positive rods (Song & Cho, 2015). A1: PYG B1: PYG Chapter 4 128 A2: FW + SS B2: FW + SS A3: FW - SS B3: FW - SS Figure 4.9: Gram stain images of E. limosum under anaerobic conditions (A) and low pressure (B). All scale bars are equal to 2 µm. 4.4.4 Cultivation and Assessments of the Subsurface Biome Growth curves were obtained of the subsurface biome by analysing the various growth studies under aerobic conditions for 32 hours (Figure 4.10), as well as under anaerobic conditions and low pressure for 48 hours (Figure 4.11) as discussed in Section 4.3.4. Chapter 4 129 100 C1: LB C2: FW + SS 10 C3: FW - SS 1 600 OD 0.1 0.01 0.001 0 4 8 12 16 20 24 28 32 Time (Hours) Figure 4.10: Growth curves of the subsurface biome under aerobic conditions (C) in orange/yellow (FW = fissure water, +SS = with sandstone, -SS = without sandstone). Very high biomass yields were obtained when the subsurface biome was grown in LB medium, but in the fissure water alone, exponential growth did not occur. When sandstone was added to the fissure water, an initial increase in the biomass can be seen, where after it immediately started to reach a plateau. This indicated that the fissure water on its own did not provide the necessary nutrients for the subsurface biome to grow optimally and that the sandstone most likely contributed some of the nutrients initially needed for growth under aerobic conditions, but could not sustain the microorganisms for exponential growth. 10 D1: LB D2: FW + SS D3: FW - SS E1: LB 605 1 E2: FW + SS OD E3: FW - SS 0.1 0 4 8 12 16 20 24 28 32 36 40 44 48 Time (Hours) Figure 4.11: Growth curves of the subsurface biome under anaerobic conditions (D) in orange/yellow and low pressure (E) in red/pink (FW = fissure water, +SS = with sandstone, -SS = without sandstone). Chapter 4 130 From the results of the anaerobic and low pressure growth studies, the best biomass yields were obtained when the subsurface biome was grown in the LB medium, without pressure. As for the rest of the growth studies, exponential growth did not occur under any of the conditions the biome was subjected to. As described in the literature, most of the deep subsurface microorganisms are living under extreme energy limiting conditions where they are likely to be inactive or to display exceptionally low metabolic activity (Jørgensen & D’Hondt, 2006; Jørgensen & Boetius, 2007). Geochemical evidence suggested that the in situ microbial metabolism in porous subsurface environments was estimated to be in the nano- to micromolar range per year, with an average turnover time for a subsurface microbial cell estimated to be decades or centuries (Phelps et al., 1994; Onstott et al., 2009). It was also found that even though subsurface microorganisms are dormant or living at extremely low metabolic rates, their diversity and persistence could offer valuable insights into life in the extremis (Phelps et al., 1994; Teske, 2005; Onstott et al., 2009). Therefore, it was not expected to obtain high biomass yields with the subsurface biome, especially when grown in the fissure water without any additional nutrients, but it was important to have a biome that could survive the various conditions and stay metabolically active to continue to contribute to carbon cycling. Additional assessments were conducted on all the growth studies using different staining and microscopy techniques (Section 3.3.6.8) and DGGE analyses (Section 4.3.6). The negative controls remained sterile. From the negative stain images (Figure 4.12), it can be seen that biomass was obtained for all the parameters tested, even though it was extremely low and that the cells were alive. The biomass from the growth studies with the fissure water contained microorganisms with varying morphology and distinctive structures, but the morphology was less diverse and more uniform for the LB medium. This suggests that when the biome was grown in the nutritionally rich LB medium, the conditions encouraged the growth or domination of selected species. As discussed in Section 3.3.6, it is very likely that the diversity within a community will change if the parameters from the culture conditions differ even slightly from the conditions of their original environment. This will introduce biases in the diversity of the community due to the methods used for isolation (Dunbar et al., 1997; Liu et al., 1997). Therefore, changes in the diversity could occur when the biome is exposed to different nutritional environments. Chapter 4 131 C1: LB C2: FW + SS C3: FW - SS D1: LB Chapter 4 132 D2: FW + SS D3: FW - SS E1: LB E2: FW + SS Chapter 4 133 E3: FW - SS Figure 4.12: Negative stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm. The live/dead stain images (Figure 4.13) indicated that the subsurface biome stayed alive under all the parameters tested. C1: LB C2: FW + SS C3: FW - SS D1: LB Chapter 4 134 D2: FW + SS D3: FW - SS E1: LB E2: FW + SS E3: FW - SS Figure 4.13: Live/dead stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm. The Gram stain images (Figure 4.14) indicated that both Gram-positive and Gram- negative microorganisms were present, again confirming the various morphological structures. Chapter 4 135 C1: LB C2: FW + SS C3: FW - SS D1: LB D2: FW + SS D3: FW - SS E1: LB E2: FW + SS Chapter 4 136 E3: FW - SS Figure 4.14: Gram stain images of the subsurface biome under aerobic conditions (C), anaerobic conditions (D), and low pressure (E). All scale bars are equal to 2 µm. 4.4.5 Microbial Diversity Assessments of the Subsurface Biome using Denaturing Gradient Gel Electrophoresis 4.4.5.1 Genomic DNA Isolation and Polymerase Chain Reaction Genomic DNA was isolated from all the growth studies with the subsurface biome (Section 3.3.6.6.1) and used to amplify the Archaeal 16S (~600 bp) (Figure 4.15B), Bacterial 16S (~1500 bp) (Figure 4.15C), and Eukaryal 18S (~1700 bp) (Figure 4.15D) rRNA gene fragments as discussed in Section 3.3.6.6.2. These experiments were done over a period of time and the agarose gels were not visualized simultaneously. Chapter 4 137 1 2 3 4 5 6 7 8 9 10 11 12 A B 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 C D Figure 4.15: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, with the amplified rRNA gene fragments for (B) TM Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRuler DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. Lanes 4 – 6 contain the LB, FW + SS, and FW - SS from the aerobic growth studies. Lanes 7 – 9 contain the LB, FW + SS, and FW - SS from the anaerobic growth studies. Lanes 10 – 12 contain the LB, FW + SS, and FW - SS from the low pressure growth studies. For the aerobic growth studies, amplicons were obtained for LB, FW + SS, and FW – SS, but only for the Bacteria domain. For the anaerobic growth studies only one amplicon was obtained, which was with LB for the Bacteria domain. For the low pressure growth studies, amplicons were obtained for LB, FW + SS, and FW – SS for all three domains. This suggests that changes in the diversity could occur when the biome is exposed to different physiological growth environments. As discussed in Chapter 1, several kinds of microbial metabolism have been described, which includes the obligate aerobes that need oxygen to grow, the obligate anaerobes that are inactivated in the presence of normal atmospheric levels of oxygen, the facultative anaerobes that can grow with or without oxygen, the microaerophiles that need oxygen to survive, but only at levels that are lower than the concentration of oxygen in the atmosphere, and the Chapter 4 138 aerotolerant microorganisms that do not need oxygen to grow, but are not inactivated in the presence thereof (Prescott et al., 1996). The diversity studies of the subsurface biome (Chapter 3) indicated that microorganisms with chemolithoautotrophic, as well as heterotrophic metabolisms are present in the fissure water, and therefore it was expected that changes in the diversity would occur in the different physiological growth environments the subsurface biome was subjected to. The bands of interest were excised from the agarose gel and purified as discussed in Section 3.3.6.6.2. 4.4.5.2 Nested Polymerase Chain Reaction The purified PCR products were used to amplify the V3/V4 hypervariable regions for Archaea (~600 bp) (Figure 4.16B), Bacteria (~600 bp) (Figure 4.16C), and Eukarya (~600 bp) (Figure 4.16D) using a nested DGGE PCR as discussed in Section 3.3.6.6.3. Amplicons were obtained for the V3/V4 hypervariable regions from all the samples. 1 2 3 4 5 6 A B 1 2 3 4 5 6 7 8 9 10 1 2 3 4 5 6 C D Figure 4.16: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons with the amplified V3/V4 hypervariable regions for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. The DGGE PCR amplicons are shown in the rest of the lanes. Chapter 4 139 4.4.5.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses The DGGE PCR products were used to construct a preliminary diversity profile for Archaea (Figure 4.17A), Bacteria (Figure 4.17B), and Eukarya (Figure 4.17C) through the use of DGGE as discussed in Section 3.3.6.6.4. 1 2 3 4 5 6 7 8 9 10 11 12 13 A B C Figure 4.17: DGGE diversity profiles for (A) Archaea, (B) Bacteria, and (C) Eukarya. Lanes 1 – 3, 8 – 10, and 11 – 13 contain the low pressure growth studies, each time in the order of LB, FW + SS, and FW – SS. Lanes 4 – 6 contain the aerobic growth studies in the order of LB, FW + SS, and FW – SS. Lane 7 contain the anaerobic growth study for LB. The DGGE profiles for the Bacteria and the Eukarya stayed mostly consistent for the fissure water, with and without the addition of the sandstone, indicating that the sandstone did not greatly influence the diversity. Growth in the fissure water with an inadequate nutrient supply has to happen under extreme energy limiting conditions and therefore, as discussed in Section 4.4.4, the microorganisms were most likely to be inactive or to display exceptionally low metabolic activity. Thus, microbial mineral dissolution from the sandstone could not have happened within the limited amount of Chapter 4 140 time the microorganisms were exposed to the sandstone during the growth studies and therefore, the potential to use the sandstone as a possible nutrient source could not occur and as a result, no profile changes in the diversity was observed. However, except for the Archaea, the profiles changed completely when grown in the LB medium. This confirmed the suggestions from the different staining images (Section 4.4.4), that the nutritional differences could influence the diversity. Even though the Bacteria were the dominant domain throughout all the parameters tested, the Archaea was the least affected by the nutritional differences in the media, as their profiles stayed fairly consistent. As discussed in Chapter 3, Bacteria is the domain that generally dominate the subsurface, with the microbial activities that dominate these extreme environments being fermentation, methanogenesis, and sulphate-reduction - the more important steps in the degradation of organics in the biogeochemical carbon cycle (Banfield et al., 1999; Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013). However, studies conducted by Morozova et al., (2010), on the composition and activity of the microbial community of a saline CO2 storage aquifer, revealed temporal out-competition of the sulphate-reducing Bacteria by the methanogenic Archaea, but the Bacteria was ultimately able to adapt to the extreme conditions of the deep subsurface and the changes under unusual conditions. This indicates that the Eukarya will most likely not be able to survive and adapt to CCS conditions, and therefore will probably not play a major role, if any, in carbon cycling in the subsurface. Sequencing and analyses were conducted on selected DGGE bands as discussed in Sections 3.3.6.6.5 and 3.3.6.6.6. After the aerobic growth studies were completed, only the Bacterial domain was found to be present, which was dominated by the Proteobacteria phylum, mainly consisting of the Alpha- and Gammaproteobacteria classes. This was followed by the Firmicutes phylum, consisting of the Bacilli class and the Actinobacteria class of the Bacteriodetes phylum. For the anaerobic growth studies in these conditions, again only the Bacterial domain was present and was dominated by the Gammaproteobacteria and the Clostridia class from the Firmicutes phylum. For the low pressure growth studies, all three domains of life were found to be present. For the Bacterial domain, the Proteobacteria prevailed as the dominating phylum, consisting of Chapter 4 141 the Alpha-, Beta-, and Gammaproteobacteria classes. This was followed by the Firmicutes phylum, consisting of the Bacilli class. Within the Archaeal domain, the main class was the Thermoprotei from the Crenarchaeota phylum. The Eukaryal domain contained the Ascomycota, the Basidiomycota, the Ochrophyta, the Alveolata, the Glomeromycota, and the Heterokontophyta. This confirmed that changes in the diversity did occur when the biome was exposed to different physiological conditions and that microorganisms with chemolithoautotrophic, as well as heterotrophic metabolisms were able to survive the various conditions. Chapter 4 142 4.5 CONCLUSIONS The positive control microorganism, E. limosum, was able to grow and produce high biomass yields in a nutritionally rich medium, under both anaerobic and low pressure conditions, with a µmax similar to that which has been reported in the literature. It was found that the introduction of low pressure did not affect the growth of E. limosum, which was expected, since this microorganism is described in the literature to be able to grow under pressures of >1 atm. High biomass yields were obtained when the subsurface biome was grown in a rich medium under aerobic conditions. However, when both E. limosum and the subsurface biome was grown in the fissure water, exponential growth did not occur, indicating that the fissure water on its own did not provide all the necessary nutrients for optimum growth. The addition of sandstone also had no effect on growth. Although exponential growth did not occur for all the parameters tested, biomass was obtained and it was confirmed that the microorganisms were still viable and metabolically active. In Chapter 3, characterization of the fissure water showed a sustained reducing environment with the presence of iron, nitrate, sulphate, and hydrogen with low levels of dissolved oxygen. Accordingly, the diversity studies of the subsurface biome indicated that microorganisms with chemolithoautotrophic and heterotrophic metabolisms are present. From this it was expected that selected species within the subsurface biome should be able to grow under aerobic and anaerobic conditions and that changes in the diversity would occur when the biome was exposed to different nutritional environments, as well as different physiological growth conditions. These changes were observed after the subsurface biome was grown under the various growth conditions and confirmed that microorganisms with chemolithoautotrophic, as well as heterotrophic metabolisms were able to survive. Care has to be taken when growth studies with subsurface biomes are done, as these biomes contain complex physiological individuals. Analyses may not always reveal the true minor community composition and often, this excludes knowledge of their specific contributions. 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Chapter 4 150 CHAPTER 5 DESIGN AND CONSTRUCTION OF THE SYRINGE INCUBATORS AND THE BIOREACTOR 5.1 INTRODUCTION It has been observed in the literature that when deep subsurface microorganisms are grown under lower or normal atmospheric pressures, their response differ from when they are grown under higher pressures, especially their metabolic activity and viability are negatively influence, since these deep subsurface biomes have adapted to their in situ high hydrostatic pressures (Yayanos & Dietz, 1983; Fang et al., 2010). Prieur, (1997), found that the optimal growth pressures for deep-sea hyperthermophiles are higher than their in situ environmental pressure. This was confirmed by Miller et al., (1988), that Methanocaldococcus jannaschii, which was isolated at 2 600 mbs (corresponding to ~260 atm, Jones et al., 1983), grew optimally at 750 atm. In contrast, Prieur, (1997), reported that deep-sea psychrophiles have optimal growth pressures below their in situ environmental pressure. It was also found that the addition of pressure, increased the maximum viable temperature of several deep-sea thermophilic Archaea (Holden & Baross, 1995; Canganella et al., 1997; Marteinsson et al., 1999a) and Boonyaratanakornkit et al., (2007), showed that under 500 atm, the optimal growth temperature for M. jannaschii was increased by 5°C, with a threefold increase in its specific growth rate. In addition, the gene and protein expression responses for thermophilic and psychrophilic piezophiles are different when grown under higher pressures. Marteinsson et al., (1999b), showed that when Thermococcus barophilus was grown at 400 atm, an unidentifiable protein was induced, while a stress protein was induced when grown under atmospheric pressure. Vezzi et al., (2005), reported that when Photobacterium profundum, isolated at 2 500 mbs, was grown under 0.1 MPa (~1 atm), genes involved in protein folding, DNA repair, and amino acid transport were activated, while at 280 atm, genes involved in respiration, fermentation and in regulating polymer degradation were activated. Furthermore, partial pressures of CO2 and H2 Chapter 5 151 above 0.01 MPa (~0.1 atm) adjusted the metabolism of several bacteria, fungi, and yeasts (Chicoye et al., 1978; Jones & Greenfield, 1982; Eklund, 1984; Lamad et al., 1988; Dixon & Kell, 1989; McIntyre & McNeil, 1998). Therefore, in order to possibly understand how these subsurface biomes will operate in situ, temperatures and pressures that characterize the deep subsurface must be mimicked using higher pressure equipment. Several experiments under various high pressures has been carried out to study the effects of high pressures on the genetic, metabolic, biochemical, and physiological aspects of microorganisms, some of which includes biohydrogenations (Schieche et al., 1997), enhanced oil and gas recovery (Bubela et al., 1987; Cloete, 2010), metabolic activities (Jannasch et al., 1996; Abe et al., 1999; Abe & Horikoshi, 2001; Bothun et al., 2004), the sterilization of microorganisms (Lin et al., 1994; Hong et al., 1997; Hauben et al., 1997; Smelt, 1998; Dillow et al., 1999; Lee et al., 2001; Zhang et al., 2006; Parton et al., 2007; Oulé et al., 2010), in situ recovery of fermentation products (Knutson et al., 1999; Berberich et al., 2000; Berberich, 2001), transcriptional responses (Boonyaratanakornkit et al., 2007), growth and survival of microorganisms under pressure (Yayanos, 1986; Nelson et al., 1991; Kato et al., 1996; Miller et al., 1988; Yanagibayashi et al., 1999; Arakawa et al., 2006; Takai et al., 2008; Hiraki et al., 2012; Sauer et al., 2012), the dissociation of ribosomes (Schulz et al., 1976), gene regulation (Bartlett et al., 1989), stabilization of proteins (Hei & Clark, 1994; Sun & Clark, 2001), and the composition of membrane lipids (Delong & Yayanos, 1985; Kaneshiro & Clark, 1995). These high pressure experiments and numerous others have all been carried out in high pressure reactors or vessels, specifically designed to fit the conditions and requirements of each experiment, some of which includes culture tubes with neoprene stoppers working as piston to pressurize the samples (Orcutt et al., 2008), glass syringes or a flexible Teflon container in a reaction chamber (Schmid et al., 1978; Hutton et al., 2001; Takai et al., 2008), flexible nickel tubes (Bernhardt et al., 1987), flexible cells made of gold (Seyfried, 1979) or titanium (Seyfried & Janecky, 1985), flexible polyvinylidene fluoride (PVDF) incubator sleeve (Sauer et al., 2012), and several types of bioreactors (Miller et al., 1988; Nelson et al., 1991; Vance & Brink, 1994; Dillow et al., 1999; Park & Clark, 2002; Bothun et al., 2004; Parton et al., 2007; Zhang et al., 2011; Hiraki et al., Chapter 5 152 2012; Sauer et al., 2012). Methods used for the addition of pressure are usually hydrostatically (Liquid only) (Zobell & Oppenheimer, 1950; Paul & Morita, 1971; Yayanos et al., 1982), hyperbarically (Both liquid and gas phases) (Sturm et al., 1987; Bernhardt et al., 1988; Miller et al., 1988; Nelson et al., 1991), or directly to solid medium (Yayanos et al., 1984). However, laboratory scale imitations of the deep subsurface conditions do have their limitations in that they may not reflect the true in situ environmental activity of the deep subsurface biomes, since the rates of microbial activity can differ orders of magnitude from what actually takes place in the subsurface (Chapelle & Lovley, 1990). Thus the equipment used in the laboratory to mimic the subsurface conditions can only be used to demonstrate the potential for the in situ microbial activity but do not give quantitative information on the processes actually taking place (Lovley & Chapelle, 1995). Therefore, in situ experiments on the reaction of the subsurface biome under CCS conditions are preferred. But, since the first pilot CO2 storage project experiment for South Africa are planned for 2017 (SACCCS, 2015), in situ environmental conditions that represents the proposed sites for CCS in South Africa (Cloete, 2010) had to be mimicked for this research, through the use of specifically designed, high pressure equipment. 5.2 AIMS OF THIS CHAPTER The main aims of this chapter were to: Design and construct a reactor(s) that can operate under higher pressures, to mimic CCS conditions within the laboratory. Chapter 5 153 5.3 MATERIALS AND METHODS Unless otherwise specified, the material used for the construction of all the syringe incubators as well as the bioreactor was 316 stainless steel. The tubing used was ¼ inch and all valves, gauges, nuts, and ferrules were from Swagelok® and designed to operate under high pressures and temperatures (maximum of ≥172 bar and 148°C). Safety precautions were used at all times while operating any of the high pressure reactors. 5.3.1 The Syringe Incubators Takai et al., (2008), have developed a technique for the cultivation and characterization of several native deep-sea chemolithoautotrophs in a gas-rich fluid under high hydrostatic pressures, corresponding to its potential in situ environments. This system is based on a liquid medium with a gas mixture inside a gas-tight glass syringe with a butyl rubber piston and sealed with a metallic needle that is closed with a butyl rubber (Figure 5.1). The syringe is then transferred to a pressure vessel, compressed with a hydraulic pump to the desired pressure and incubated (Figure 5.2). By using this system, Takai et al., (2008), were able to successfully study the growth, survival, and methane production of Methanopyrus kandleri strain 116, under high temperatures and hydrostatic pressures. Figure 5.1: Schematic illustration of the microbiological cultivation and incubation technique (Taken from Takai et al., 2008). Chapter 5 154 Figure 5.2: Procedure for cultivation in a gas-rich fluid under high hydrostatic pressures (Taken from Takai et al., 2008). Hutton et al., (2001), have used a similar system where sealed syringes were incubated inside a pressurized vessel (Figure 5.3) to test the effect of hydrostatic pressure on disc cells. Figure 5.3: Schematic illustration of the syringes inside the pressure vessel (Taken from Hutton et al., 2001). Chapter 5 155 Cultivation of microorganisms in this manner has the advantage that the numerically predominant or the functionally significant microbial populations in the in situ environments can be characterized because the in situ hydrostatic pressures is a potential key physical parameter which directly affects the growth of the microorganisms (Abe et al., 1999; Kato, 2006; Takai et al., 2009). But more importantly, the pressures and temperatures are the main physicochemical factors preparing the thermodynamic states of energy and carbon metabolisms for microbial growth. This means that the available gaseous substrates that are dissolved in the liquid phase of the growth medium and much less abundant under lower or normal atmospheric pressures than when compared with the pressures in their in situ environments (Charlou et al., 2002; Person et al., 2005; Lupton et al., 2008; Takai et al., 2009). If these in situ environmental parameters are ignored in laboratory experiments, metabolic biases can be introduced, due to the enrichment of certain populations that are less dominant in the in situ environments but are energetically favourable in laboratory experiments (Takai et al., 2009). Therefore, the same cultivation technique [mainly from Takai et al., (2008)] was used to study the potential growth and survival of the subsurface biome under high pressures and in a gas-rich environment. 5.3.1.1 Design and Construction All schematic illustrations for the syringe incubators (Figure 5.4) were created with the solid modelling, computer-aided design (CAD) and computer-aided engineering (CAE) software program, SolidWorks®. The schematic illustrations are not according to scale. Each incubator consisted of a hollowed stainless steel vessel that was fitted with a Teflon syringe ring (Figure 5.4B and E). A syringe was inserted through the hole in the middle of the ring and the holes on the outside of the ring allowed circulation of the water inside the vessels. The lid of the incubator (Figure 5.4C) was made to fit tightly into the vessel and therefore pressure was needed to insert the lid into the vessel. Thus, a handle bar was fitted through the lid which was used to apply the pressure. To ensure a completely sealed-off incubator, a double seal was added, consisting of two high- pressure Neoprene O-rings with Teflon spacers. As a safeguard mechanism to ensure that the lid remained in place while under pressure, a safety pin (Figure 5.4D) was Chapter 5 156 inserted all the way through the vessel and the lid (Figure 5.4A). An opening was made through the lid (Figure 5.4B) for pressurization of the incubator. A B C D E Figure 5.4: Schematic illustrations of (A) the complete syringe incubator, (B) a cut view of the syringe incubator, (C) the lid, (D) the safety pin, and (E) the syringe ring. A needle valve and pressure gauge (Figure 5.5) was attached to the lid. In this way the syringe incubators could be pressurized to the desired pressure using either gas or a hydraulic pump. These incubators were designed for either 5 ml or 25 ml syringes. Chapter 5 157 Except for the Swagelok® components, everything on the syringe incubators was custom-made by the Department of Instrumentation at the UFS (Bloemfontein, RSA). C A D E B F Figure 5.5: Images of (A) all the constructed syringe incubators, (B) the individual components of a syringe incubator, (C) the lid, (D) the 5 ml syringe ring, (E) the 25 ml syringe ring, and (F) the safety pin. Chapter 5 158 5.3.1.2 Syringe Preparation All syringe preparations were conducted under sterile conditions in a laminar flow cabinet (Labotec®). The syringes used are non-autoclavable, gas-tight, glass syringes (Hamilton), fitted with 22 gauge stainless steel needles (Hamilton) and sealed with blue chlorobutyl rubber stoppers (Bellco Glass Inc.). Before and after each usage, the syringes, needles and rubber stoppers were sterilized with 70% EtOH, followed by rinsing three times with sterile dH2O. The desired amount of inoculated medium was added to each syringe, followed by the insertion of the specific gas mixture. The gas mixture was prepared as discussed in Section 4.3.3, but in empty, sterile, serum vials that were crimp sealed with a blue chlorobutyl rubber stopper and an aluminium cap. The gas was transferred to the syringes by inserting the needle that is attached to the syringe, into the serum vial. The syringe was sealed with a blue chlorobutyl rubber stopper, immediately after the gas was added. 5.3.1.3 Operational Procedure The syringe incubators were filled with water and heated to the operating temperature, 24 hours before use, through incubation in a temperature-controlled dry oven (Therm-O- Mat). This ensured that the stainless steel was already at the desired temperature and therefore the pressure stayed relatively consistent when incubated after pressurization. The water inside the reactors ensured that the internal syringe pressure is in equilibrium with the external pressure. The prepared syringes were transferred into the heated incubators and the incubators were sealed and secured with the safety pin. A hydraulic pump (Jackco®) was connected to the incubators and pressurized to the desired pressure (Figure 5.6). While the pressure was added, the incubators were submerged in a thermostatic water bath [Gesellschaft für Labortechnik (GFL®)], pre-heated to the operating temperature. The needle valve was closed and the incubators were transferred to the temperature-controlled dry oven for incubation. The hydraulic pump was modified so that water was used to pressurize the incubators instead of oil. This was done by connecting the hydraulic pump to a cylinder which was divided in the middle with a piston system. The one side of the cylinder was filled with oil and the other side with water. The hydraulic pump was connected to the oil cylinder and the incubators to the water cylinder. The pump releases oil into the oil cylinder which allows the piston to push Chapter 5 159 down on the water cylinder. Water is then expelled from the cylinder to pressurize the incubators. When the water cylinder was empty, it was connected to a tap and filled with water. Temperature regulation of the incubators was very important throughout the pressurization and operation steps to ensure that the desired pressure was maintained as closely as possible. A B Figure 5.6: (A) Pressurizing the syringe incubators using a hydraulic pump and (B) the modified cylinder connected to the hydraulic pump. The white tubing is used for filling the water cylinder. Chapter 5 160 To depressurize the syringe incubators, the needle valves were simply opened slowly and the culture from the syringes was added to sterile falcon tubes to be used for analyses in further experiments. 5.3.2 The Bioreactor With the syringe incubators, the experiments were considered as batch studies. These experiments were done without the addition of sandstone and each syringe was sacrificed at certain time intervals displaying the reactivity of the biome for this period. Care was taken that all syringes in a set of batch experiments contained the same amount of cells. Consequently, no circulation or continuous monitoring was possible. It was also difficult to accurately control the pressures, since the smallest change in temperature during either the pressurization or the incubation resulted in a substantial change in the pressure. In order to study the subsurface biome as a continuous culturing experiment, a novel bioreactor was designed to operate under high pressures in the presence of the supercritical fluid. Even though it was not needed for this research project, the bioreactor is also able to operate at high temperatures. 5.3.2.1 Designs and Constructions All schematic illustrations for the bioreactor were created using the software program, AutoCAD. This bioreactor was adapted several times to improve its safety and operation and only when major differences were made, it has been indicated through schematic illustrations. In all designs, the CO2 was used to saturate the reactor and to create the initial pressure. The desired operating pressure and temperature was always ≥80 bar and 35°C, respectively. The schematic illustrations are not according to scale. The first bioreactor (Figure 5.7) was designed to be a flow-through system, with a constant supply of fresh medium via a high-performance liquid chromatography (HPLC) pump (Shimadzu), but the reactor could be isolated from the medium supply, when needed. The reactor itself consisted of a 316 stainless steel pipe and has an inner diameter of 2.1 cm and 24.5 cm in length (Volume of 84.86 cm3). Both ends of the pipe body are fitted with high pressure nuts and ferrules. Chapter 5 161 Figure 5.7: Schematic illustration of the first design of the bioreactor. Over time, small adjustments were made to this design, which included adding a port in the media reservoir for the insertion of additional medium or electron donors when needed. A non-return valve was added before the reactor, to ensure that the gas and medium can only flow through the reactor in one direction and a pressure release valve was added to the system as a safety measurement to prevent pressure build-up. A pressure gauge was added before the reactor to measure the pressure inside the reactor and a liquid sampling port was added at the outlet of the reactor, with a gas sampling port before the reactor. With this bioreactor, the kinetics and continuous measurements could not be done, since fresh medium would have been supplied at all times, therefore no circulation of the liquid would take place. Contact time with the biome was also limited and if the reactor would have been isolated from the media supply, the system would Chapter 5 162 again be a batch experiment as with the syringe incubators. Therefore, no usable data could be obtained. For the second design (Figure 5.8), the system was changed to allow circulation of the liquid using the HPLC pump and the CO2 gas was used to introduce the medium into the reactor. A sieve was inserted into the reactor that served as a filter to prevent blockage of the HPLC pump. Isolating valves were added at strategic points to ensure that the reactor could be closed off without losing pressure. Figure 5.8: Schematic illustration of the second design of the bioreactor. Chapter 5 163 Final adjustments to this design included to regulate the pressure more effectively and were added to the next design (Figure 5.9) only had a few practical changes which included the addition of a pressure gauge before the media reservoir to ensure that the pressure created with the gas was higher than the pressure inside the reactor for the media supply. An additional pressure release valve was added before the media reservoir and the sieve was removed from the reactor and replaced with an in-line pre- filter after the reactor to ensure that the filter can be removed or changed, if needed, without having to open or disturb the reactor. Figure 5.9: Schematic illustration of the third design of the bioreactor. Chapter 5 164 This bioreactor was constructed (Figure 5.10) and mounted inside a safety cabinet for protection. Only the HPLC pump and the CO2 gas cylinder were outside the safety cabinet. A heating-coil was wrapped around the reactor and the media reservoir to regulate the temperature. REACTOR LIQUID SAMPLING PORT MEDIA RESERVOIR HPLC SECURITY GUARD PRESSURE RELEASE VALVES GAS SAMPLING PORT HEATING-COIL A B HPLC PUMP SAFETY LINE TO THE CABINET CO2 GAS C Figure 5.10: (A) The constructed bioreactor, (B) the bioreactor inside the safety cabinet, and (C) the HPLC pump connected to the bioreactor, with the line to the CO2 gas cylinder, visible outside of the safety cabinet. Unfortunately, with this design, a rapid drop in pressure with large increments were obtained, especially during each sampling, thus resulting in the CO2 not being in the supercritical condition. A pressure shock on the microorganisms that were introduced each time a sample was withdrawn had to be considered. Also, the system would have Chapter 5 165 been under subcritical CO2 conditions for extended periods of time, since the HPLC pump was not able to restore the pressure rapidly enough. The HPLC pump was also able to maintain the pressure for only short periods of time, since the inlet of the pump needs to be under low pressure and therefore, the pump was not able to handle the system pressure (≥80 bar) on the inlet. The heating-coil around the reactor and media reservoir alone was also not sufficient enough to regulate the temperature. For the next design (Figure 5.11 and Figure 5.12), the media reservoir was replaced to a variable media reservoir, attached to a piston pressure regulator. In this design, the CO2 was no longer used to introduce the medium into the reactor, but through the piston pressure regulator connected to a hydraulic pump (Jackco®). This piston regulator was connected to a pressure gauge, to ensure that it is always under the desired pressure and circulation still occurred through the HPLC pump. However, an adjustable pressure valve was included in the system before the HPLC pump. This valve was set to open at the operating pressure to allow small amounts of liquid to pass through each time the valve opened. This meant that everything before the adjustable pressure valve was under the high operating pressure and between the valve and the HPLC pump the liquid could be maintained under atmospheric pressure. After the HPLC pump, the high operating pressure was restored. Several techniques, described in the literature, are used to obtain the desired temperature in the high pressure reactors, including thermistors (Bernhardt et al., 1987), water baths (Jannasch et al., 1996), and drying ovens/incubators (Miller et al., 1988; Takai et al., 2008). For this design, the heating-coil was replaced with a temperature controlled fan inside the safety cabinet and connected to an uninterruptible power supply (UPS), converting the safety cabinet into an incubator. All the openings in the safety cabinet were sealed and the temperature was verified with a thermometer. The idea behind this system was that each time a sample was collected, the pressurized piston regulator immediately thrusts down on the variable media reservoir to keep the system under the correct pressure. The hydraulic pump was then used to restore the pressure in the piston pressure regulator. This piston regulator was designed to ensure that the oil from the pump only enters the piston regulator and never come in contact with the reactor. However, to achieve this, the collection of the sample and the Chapter 5 166 pressurizing of the piston pressure regulator had to occur simultaneously. Even though the two actions were manually carried out at the same time, the drop in pressure during sampling was faster than restoring the pressure with the hydraulic pump. This resulted in a double pressure shock on the microorganisms, once during sampling and again during re-establishing of the pressure. The HPLC pump was only able to maintain the pressure for short periods of time, especially since gas bubble which entered the pump, prevented it from operating continuously. Gas bubbles formed each time when the pressure changed through the adjustable pressure valve to atmospheric pressure. To compensate, a small reservoir using a sterile serum vial, was included as a gas trap, just before the HPLC pump (Figure 5.12). However, a head space formed inside the reservoir from all the gas in the system. This increased the pressure on the small reservoir until it was too high for the inlet of the HPLC pump to handle. If the serum vial was to be replaced each time the pressure was too high, it would have meant that each time the reservoir was removed, a certain amount of the gas inside the system were removed as well. Sauer et al., (2012), included an empty HPLC column between their reactor and HPLC pump which served as a gas trap to prevent the entry of gas bubbles into the pump. They mounted the column vertically and the medium was flowing from the top to the bottom, trapping the gas bubbles at the top. Chapter 5 167 Figure 5.11: Schematic illustration of the fourth design of the bioreactor. Chapter 5 168 LINE FROM THE TEMPERATURE HYDRAULIC PUMP CONTROLLED INTO THE PISTON FAN REGULATOR PISTON PRESSURE REGULATOR REACTOR UPS SAMPLING PORT LINE TO THE CO2 GAS VARIABLE MEDIA RESERVOIR HYDRAULIC PUMP HPLC PUMP PISTON PRESSURE REGULATOR FROM PUMP OUTLET INTO THE MEDIA GAS RESERVOIR TRAP FROM REACTOR TO PUMP INLET VIA THE GAS TRAP ADJUSTABLE PRESSURE VALVES THERMOMETER VARIABLE MEDIA RESERVOIR Figure 5.12: The bioreactor, constructed from the fourth design. Chapter 5 169 Even though a design like the one from Sauer et al., (2012), could have possibly solved the problem, it was decided to completely remove the HPLC pump from the system in the fifth design (Figure 5.13 and Figure 5.15). Thus, to ensure circulation of the liquid through the system, a second variable media reservoir, attached to a piston pressure regulator, was included. The double piston regulators worked in opposite directions via three-way valves and operated similar to a syringe pump. The hydraulic pump from the fourth design was replaced with nitrogen gas. The advantage in using the gas is that the pressure is immediately restored after a sample has been withdrawn, since the nitrogen gas cylinder was kept open at all times and the regulator was set to the desired pressure. Several non-return valves were included in the system to ensure that the circulation always occurred in one direction. This system should ensure that the liquid, from the media reservoir, was inserted into the system with a peristaltic pump and sterile Masterflex® Norprene tubing (Cole-Parmer®). Due to the presence of non-return valves, the liquid entered the system from below the reactor until the entire system, including both the variable media reservoirs, were filled with the media. The CO2 gas was then used to saturate the system, also from below the reactor, and to give the initial pressure (~55 bar). Since the CO2 gas cylinders are not under enough pressure to reach ≥80 bar, the rest of the pressure was added with the piston regulators and nitrogen gas. To reach the desired pressure, the first piston pressure regulator was pressurized, while the second piston pressure regulator was set to release pressure via the pressure release valve. This meant that the three-way valve from the first piston regulator was opened between the nitrogen gas cylinder and the piston regulator which allowed the nitrogen to enter and adjust the first variable media reservoir, while the three-way valve from the second piston regulator was opened between the piston regulator and the pressure release valve to allow the nitrogen inside the piston regulator to escape. The same principle was applied to the variable media reservoirs through the opening of the three-way valves, allowing the first media reservoir to empty its contents into the system using the piston pressure regulator, while the second media reservoir was filled with liquid from the system, pushing up on the piston pressure regulator. The moment the first media reservoir was empty, all four the three-way valves were switched to ensure that the same process was repeated just in the opposite direction. Chapter 5 170 Even though the flow direction was altered between the two media reservoirs, the flow through the reactor was always in the same direction and was controlled with the addition of the non-return valves. The flow speed of the system was controlled by opening and closing the pressure release valve until the desired flow was reached. As an additional control, a control valve was included in the system and isolating valves were added throughout the system to ensure that any part of the system can be removed, if faulty, without disturbing the pressure of the reactor itself. Pressure gauges were connected to the reactor, the variable media reservoirs and the piston pressure regulators to ensure that the entire system was under the desired pressure at all times. As a safety measurement, an adjustable pressure valve was included in the system which was always set to release at a pressure of 5 bar higher than the operating pressure. To enable sampling, a liquid sampling port was connected to the adjustable pressure valve to allow samples to be taken in a controlled manner. The piston regulator that was under pressure, ensured that the operating pressure was maintained by decreasing the system volume after sampling. Chapter 5 171 Figure 5.13: Schematic illustration of the fifth design of the bioreactor. Chapter 5 172 For the final design (Figure 5.14 and Figure 5.15), small adjustments were made, but the mode of operation stayed the same as discussed in design five. The filter added after the reactor was removed, since blockage occurred. The flow speed was still too fast, even though it was regulated with the control valve and the pressure release valve. Therefore, a flow restrictor was included in the system using 1/16 inch stainless steel tubing. This was added after the adjustable pressure valve with the sampling port and was connected via a three-way valve to ensure that the flow restrictor could be isolated when it was not needed. A liquid inlet port was added between the two variable media reservoirs to allow for any additional liquid to be introduced into the system by isolating one of the media reservoirs and releasing its pressure. The liquid can then be introduced into the media reservoir at atmospheric pressure. The media reservoir was pressurized again before opening to the system, thus the pressure on the reactor itself was never disturbed. Both the nitrogen and the CO2 gas lines were equipped with valves that isolated them and could be used to evacuate or purge the lines to allow them to be removed from the system without being under pressure. All gas cylinders were connected to the bioreactor using high pressure, safety tubing. The bioreactor was designed to be able to operate under high pressures and temperatures and can be used for both static and circulating experiments. The bioreactor had a total system volume of ~400 ml. Except for the Swagelok® components and all the pumps used, everything on the bioreactors was custom-made by the Department of Instrumentation at the UFS (Bloemfontein, RSA), the Department of Electronics at the UFS (Bloemfontein, RSA) and Earthmoving Repair Services (Bloemfontein, RSA). Chapter 5 173 Figure 5.14: Schematic illustration of the sixth design of the bioreactor. Chapter 5 174 NON-RETURN VALVES GAS TIGHT VALVE D E CONTROL VALVE TO CO2 CYLINDER A B TO PERISTALTIC C PUMP F PRESSURE RELEASE TEMPERATURE VALVE CONTROLLED FAN UPS NITROGEN CYLINDER CO2 CYLINDER MEDIA RESERVOIR PERISTALTIC PUMP G Figure 5.15: (A) The bioreactor, constructed from the fifth and sixth designs, (B) a gas tight syringe connected to the sampling port, (C) the reactor, (D) the three-way valves regulating the variable media reservoirs, (E) the three-way valves regulating the piston pressure regulators, (F) the connection to the CO2 gas cylinder and the media reservoir through the peristaltic pump, and (G) the bioreactor as a whole. Chapter 5 175 5.3.2.2 Operational Procedures All connections were secured to ensure a leak free system. The bioreactor was then washed overnight with 10% (v/v) HCl to remove any organic contaminants and impurities (Rotzsche, 1991), by slow circulation through the entire system. This was followed by an overnight sterilizing step, by slowly circulating 10% (v/v) commercial bleach through the system. The system was then rinsed with Nano-pure water, until all traces of the bleach was removed. The reactor was removed from the system and packed with the rock matrix under sterile conditions in a laminar flow cabinet (Labotec®), where after the reactor was again connected to the system. The temperature controlled fan was switched on and the system was allowed to reach the operating temperature, where after the desired medium, containing the inoculum was added to the reactor until the entire system was filled using a peristaltic pump and sterile Masterflex® Norprene tubing (Cole-Parmer®). This liquid was added by bypassing the flow restrictor. CO2 gas was then used to saturate the system and to add the initial pressure, by opening the regulator in the CO2 gas cylinder to its maximum. The piston pressure regulators were then pressurized with the nitrogen gas, by setting the regulator on the nitrogen gas cylinder to the desired operating pressure. The three-way valves were opened to allow circulation through the flow restrictor. The CO2 gas line was removed from the system, as well as the line used to fill the reactor from the media reservoir. Whenever fresh media had to be introduced into the system, the latter line was attached to an additional liquid inlet port between the two variable media reservoirs. One of the media reservoirs was isolated from the system and evacuated to atmospheric pressure. The fresh liquid was added into the media reservoir using the peristaltic pump, where after the media reservoir was pressurized and opened to be part of the system. To obtain a sample, a sterilized, gas-tight glass syringe (Hamilton) was connected to the sampling port via a gas-tight valve. The pressure valve was opened to allow liquid to slowly enter into the syringe, where after the gas tight valve connected to the syringe was closed to ensure no gas escapes from the syringe. In this way, samples were taken in a controlled manner without losing system pressure or altering any of the conditions. The syringe was removed from the system and the collected sample was injected into a sterile, gas-tight vial. Chapter 5 176 To end the run, all valves in the system were closed to isolate the reactor, followed by closing the nitrogen gas cylinder. The pressure on the system was slowly released by opening the pressure release valve of the piston regulators. This was done for both the variable media reservoirs. The liquid was pumped from the system using the peristaltic pump and collected in sterile containers to be used for analyses. The reactor was removed from the system and opened under sterile conditions, where the rock was removed for analyses. Throughout preparation and operation of the bioreactor, leak tests were constantly done to ensure a system free from any gas leakages. The final pressure in the system was always controlled with the regulator on the nitrogen gas cylinder and the flow speed was regulated by connecting a tube to the pressure release valve from the piston regulators. This tube was inserted into water to allow the visualization of bubbles that formed as the nitrogen gas was released. The pressure release valve was adjusted to obtain the desired flow speed. Chapter 5 177 5.4 CONCLUSIONS Various authors have reported on different systems which have been used to study the effects of higher pressures on the genetic, metabolic, biochemical, and physiological aspects of microorganisms and these systems have been designed to fit the specific conditions and requirements of each experiment. The reactors used in this research were successfully designed to be versatile and corrosion resistant and were able to operate under higher pressures and temperatures, to mimic CCS conditions within the laboratory. Sampling from the bioreactor was possible, without disturbing any of the reactor conditions. Even though the bioreactor functioned very well after the sixth design, enhancements can still be made to improve the functionality of the bioreactor. Since the three-way valves had to be switched manually each time one of the variable media reservoirs were empty, it meant that the bioreactor had to be monitored continuously to ensure constant circulation. To prevent this, one might consider the use of automatic valves for the switching. The material used in constructing the reactors were made out of corrosion-free stainless steel, but any potential corrosion and metal leakage from the bioreactor should be considered. Thus, Nelson et al., (1992), had the interior of their reactor plated with 24- karat gold. To prevent the gold coating from flaking and peeling off under high pressures and mechanical stresses, they first added an undercoat of electroless nickel below the gold. This enabled better adhesion of the gold and increased the mechanical strength of the coating. Sauer et al., (2012), only used inert materials such as polyvinylidene fluoride (PVDF), perfluoro-elastomers (FFKM), or gold during the construction of their bioreactor, to prevent corrosion. On another note, the design only allowed for a total reactor volume of 84.86 cm3 which was a very limited sample to be used for comprehensive analyses and should therefore be considered to adapt the reactor to bigger volumes. Chapter 5 178 5.5 REFERENCES Abe, F. and Horikoshi, K. (2001). The Biotechnological Potential of Piezophiles. Trends Biotechnol. 19:102-108. Abe, F., Kato, C. and Horikoshi, K. (1999). 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Zhang, J., Davis, T.A., Matthews M.A., Drews, M.J., LaBerge, M. and An, Y.H. (2006). Sterilization using High-Pressure Carbon Dioxide. Journal of Supercritical Fluids. 38:354-372. Zhang, Y., Maignien, L., Zhao, X., Wang, F. and Boon, N. (2011). Enrichment of a Microbial Community Performing Anaerobic Oxidation of Methane in a Continuous High-Pressure Bioreactor. BMC Microbiology. 11:137. Zobell, C.E. and Oppenheimer, C.H. (1950). Some Effects of Hydrostatic Pressure on the Multiplication and Morphology of Marine Bacteria. J. Bacteriol. 60:771-781. Chapter 5 187 CHAPTER 6 HIGH PRESSURE STUDIES USING THE SYRINGE INCUBATORS 6.1 INTRODUCTION In Chapter 4 it was shown that the subsurface biome was able to stay viable and metabolically active after being subjected to various environmental changes, which included nutrient deprivation and relatively low pressure. However, as discussed in Chapter 1, geological carbon sequestration would require storage at depths of ~800 mbs which are associated with higher pressures and temperatures, thereby causing the CO2 to be in a supercritical state (Benson & Cole, 2008; Cunningham et al., 2009; Cloete, 2010; Nondorf et al., 2011). Thus, like Mu et al., (2014), and other authors, an important question to address during CCS is how will the subsurface microbial biosphere respond to increased levels of CO2? An increase in the depth of the terrestrial subsurface results in even bigger environmental changes such as an increase in the pressure, temperature, and salinity of the water, and a possible decrease in the redox potential, pore space, and nutrient abundance (Machel & Foght, 2013; Wilkins et al., 2014). As a result, the microorganisms living in these environments have to be able to survive and multiply under these conditions (Wilkins et al., 2014), thus, living under extreme energy limiting conditions under which the microorganisms are likely to be inactive or to display exceptionally low metabolic activities (Phelps et al., 1994; Jørgensen & D’Hondt, 2006; Jørgensen & Boetius, 2007; Onstott et al., 2009). Therefore, the subsurface biome was subjected to different conditions which the microorganisms will encounter during CCS, including high pressures and high CO2 concentrations in both nutrient rich and nutrient deprived environments. Chapter 6 188 6.1.1 Supercritical Carbon Dioxide At standard temperature and pressure (STP), CO2 will behave as a gas in air, or as a solid (dry ice) when frozen. If both the temperature and pressure are increased from STP to be at or above the critical point for CO2 (73.8 bar and 31.1°C) (Figure 6.1), it will be in a supercritical state where the CO2 has a gas-like viscosity and the density of a liquid, but it is neither a gas nor a liquid and can therefore only be defined as a substance in a state (supercritical) above its critical temperature (Tc) and critical pressure (Pc) (Dostal et al., 2004; Garcia-Gonzalez et al., 2007; Mitchell et al., 2009; Cloete, 2010; Budisa & Schulze-Makuch, 2014). If CO2 is stored at depths of ~800 mbs, the higher pressures and temperatures at these depths will result in the CO2 being in a supercritical state and therefore, able to fill the pore spaces more efficiently, as well as decreasing the buoyancy differences when compared with in situ fluids (Benson & Cole, 2008). In the supercritical state, CO2 can also dissolve more readily in water and react with cations to form stable mineral compounds (Metz et al., 2005; Cloete, 2010). At depths less than 800 mbs, where the pressures and temperatures are lower, the CO2 will be in a subcritical state (liquid form), where it will be less soluble and not form a homogeneous mixture when in contact with the formation water, therefore, reactions where stable, mineral compounds are created, will be less (Metz et al., 2005; Cloete, 2010). Figure 6.1: A pressure-temperature phase diagram for carbon dioxide (Taken from https://en.wikipedia.org/wiki/Supercritical_carbon_dioxide). Chapter 6 189 However, during the injection of supercritical CO2 (sc-CO2), the pressure in the area surrounding the injection point will increase, creating a possible upward hydrodynamic pressure gradient that can progress through the cap-rock that is responsible for trapping the CO2 in the storage rock (Chapter 1). This can result in possible upward leakage of CO2 through fractures near the injection wells (White et al., 2003; Mitchell et al., 2009), allowing for enhanced interaction between sc-CO2 and the subsurface biomes. 6.1.2 Microbial Interaction with Supercritical Carbon Dioxide The distinctive properties of sc-CO2 under high pressures have led to it being used in a variety of fields such as the food and beverage, pharmaceutical, and biomedical industries, where it is often used as a sterilizing agent because it is non-toxic, non- flammable, and does not degrade polymers or heat-sensitive materials (Chen & Ling, 2000; Rozzi & Singh, 2002; Zhang et al., 2006; Glossner, 2013; Peet et al., 2015). It has been found that the injection of sc-CO2 into the deep subsurface affects the majority of microorganisms negatively, mostly due to its low viscosity and low surface tension. This results in direct physiological impacts on the subsurface microorganisms because it can quickly penetrate complex cellular material to reduce the number of viable cells (McHugh & Krukonis, 1994; Zhang et al., 2006; Mitchell et al., 2009; Basava- Reddi, 2012; Peet et al., 2015). This microbiocidal effect is enhanced at higher temperatures which will increase the fluidity of cellular membranes, leading to enhanced CO2 penetration through the cell walls, as well as at higher pressures which will increase the diffusivity and solubility of sc-CO2 (Lin et al., 1994; Hong et al., 1997; Mitchell et al., 2009; Basava-Reddi, 2012). As described in the literature, the mechanism for cell deactivation through the use of sc- CO2 under high pressures contains several steps that can take place simultaneously and in a very complex and organized manner. These steps include: 1.) solubilization of pressurized CO2 in the external liquid phase, 2.) modification or solubilization of the cellular membranes, 3.) cell lysis due to depressurization, 4.) decreasing of the intracellular pH (pHi), 5.) inactivation of the key enzymes or inhibition of the cellular metabolism due to the lowering of the pHi, 6.) direct inhibitory effects of molecular CO2 Chapter 6 190 − and HCO3 on metabolism, 7.) disordering of the intracellular electrolyte balance, 8.) extraction of vital components from cells and cell membranes, and 8.) interfering with protein synthesis (Garcia-Gonzalez et al., 2007; Basava-Reddi, 2012; Li et al., 2013). Therefore, the longevity of geological carbon sequestration does not only depend on the trapping mechanisms (solubility, mineral, and physical), but also on the interaction between the subsurface microorganisms that contribute to biogeochemical cycling in the injection formations, and the sequestered CO2 (Glossner, 2013). 6.2 AIMS OF THIS CHAPTER The main aims of this chapter were to: Evaluate the subsurface biome from the Star Diamonds mine in an environment that mimics geological CCS. Study the potential survival and possible growth of the subsurface biome under high pressures and high CO2 concentrations in both nutrient rich and nutrient deprived environments. Chapter 6 191 6.3 MATERIALS AND METHODS For all the high pressure experiments throughout this chapter, the syringe incubators, as described in Section 5.3.1, were used. The syringes used for all experiments were sterilized and prepared as discussed in Section 5.3.1.2. All graphs were constructed by plotting the mean and error values from two independent studies. 6.3.1 Cultivation Conditions and Analyses for Eubacterium limosum To study the growth of E. limosum under various pressures, PYG medium was prepared anaerobically using fissure water as discussed in Section 4.3.1. The 5 ml syringes were prepared by adding 2 ml of the PYG medium containing a 10% (v/v) inoculum of E. limosum (Figure 6.2A), followed by adding 3 ml of a gas mixture, containing 80% H2 and 20% CO2 (v/v) (Figure 6.2B) as discussed in Section 4.3.3. The 25 ml syringes were prepared by adding 4 ml of the PYG medium containing the 10% inoculum (Figure 6.2C), followed by adding 6 ml of the gas mixture (Figure 6.2D). A B C D Figure 6.2: The prepared syringes for (A) 5 ml without gas, (B) 5 ml with gas, (C) 25 ml without gas, and (D) 25 ml with gas. (The LB medium is displayed in the yellow colour and the PYG medium in the red). Chapter 6 192 The syringes were transferred into heated incubators and pressurized as discussed in Section 5.3.1.3, until the desired pressure was reached. The pressurized syringes were then incubated at 35°C for 48 hours where after it was depressurized and the culture was analysed. For each syringe that was incubated for 48 hours, a duplicate set was prepared and pressurized, but immediately depressurized again. This was used as the time zero analyses. Negative controls, containing no cells, were also prepared for each pressure tested. The pressure range tested was from 10 – 100 bar, with 10 bar increments. Final growth yields were analysed after each syringe was sacrificed, by measuring the OD at 600 nm using a Spectronic® GenesysTM 5 spectrophotometer (Spectronic Instruments). Adenosine triphosphate (ATP) measurements were done using the CellTiter-Glo® Luminescent Cell Viability Assay (Promega) according to the manufacturer’s instructions, with a Glomax® Multi+ Detection System (Promega). For additional confirmation of microorganism viability, the LIVE/DEAD® BacLightTM Bacterial Viability Kit (Molecular Probes, Inc., Invitrogen, Promega) was used as discussed in Section 3.3.6.8.2. To study the growth of E. limosum under various gas concentrations, syringes were prepared with the PYG medium containing a 10% (v/v) inoculum of E. limosum. The desired gas mixtures were prepared using the procedure explained in Section 4.3.3 and added to each syringe. The syringes were transferred to the prepared incubators and pressurized to 80 bar, to ensure that the CO2 is in the supercritical state. For each syringe, a duplicate set was prepared that was pressurized to only 70 bar, to ensure that the CO2 is in the subcritical state. In this way, the influence of supercritical CO2 could be evaluated under the same conditions as subcritical CO2. The pressurized syringes were incubated and analysed as described previously. Time zero analyses and negative controls were again prepared for each test. Over a period of time, care was taken to consider any temperature fluxes, thus the incubators were constantly monitored to ensure growth took place under the correct pressure. The gas range tested were different combinations of a H2:CO2 ratio and included 80%:20% (v/v), 50%:50% (v/v), 20%:80% (v/v), and 0%:100% (v/v). Chapter 6 193 To study the growth of E. limosum in both nutrient rich and nutrient deprived environments, syringes were again prepared with the desired medium containing a 10% (v/v) inoculum of E. limosum. Media used were prepared anaerobically and included -1 -1 PYG (Section 4.3.1), FW (Section 4.3.3), a minimal medium (0.2 g.L KH2PO4, 0.25 g.L -1 -1 -1 -1 -1 NH4Cl, 1 g.L NaCl, 0.4 g.L MgCl2, 0.5 g.L KCl, 0.1 g.L CaCl2, 0.2 g.L MOPS, 0.1 g.L-1 yeast extract, 1 ml.L-1 vitamin solution, 1 ml.L-1 trace element solution, pH 7) with the addition of glucose (2 g.L-1) (MM + Glc), and a minimal medium without the addition -1 -1 of glucose (MM - Glc). All media contained 0.5 g.L cysteine-HCl.H2O and 1 mg.L resazurin and the PYG and MM were prepared using fissure water. The gas, containing 0% H2 and 100% CO2 (v/v), was added as described previously and the syringes were pressurized to 80 bar, for supercritical CO2 conditions and to 70 bar, for subcritical CO2 conditions. The pressurized syringes were incubated and analysed as described previously and time zero analyses and negative controls were again prepared for each test. 6.3.2 Cultivation Conditions and Analyses for the Subsurface Biome The growth of the subsurface biome was studied using the same pressures, gas concentrations and media compositions that were used for the cultivation of E. limosum as discussed in Section 4.3.3. The only differences were that general LB medium (Section 4.3.4) was used and not the PYG medium and the 10% (v/v) inoculums were made using the fissure water, containing the TFF concentrated subsurface biome (Section 3.3.6.3). Time zero analyses and negative controls were prepared for each parameter and the cultures from all the syringes were analysed for growth and viability as discussed in Section 4.3.3. Chapter 6 194 6.3.3 Experimental Layout for the Cultivation of Eubacterium limosum and the Subsurface Biome A B Figure 6.3: A summary of the experimental layout for the cultivation of E. limosum and the subsurface biome under increasing pressure and (A) different gas concentrations, with (B) different media compositions. Chapter 6 195 6.4 RESULTS AND DISCUSSIONS 6.4.1 Cultivation and Analyses of Eubacterium limosum Optical density was used to analyse the growth of E. limosum after sacrificing the syringe experiments incubated under different pressures (Figure 6.4), different CO2 concentrations (Figure 6.5), and different media compositions (Figure 6.6) as discussed in Section 4.3.3. Figure 6.4: The growth of E. limosum after 48 hours in PYG medium, under various high pressures and 20% CO2. The increase in biomass yield obtained for E. limosum after 48 hours of growth indicated that the ability of E. limosum to grow under pressure was not influenced by the increasing pressure, even up to 100 bar. Figure 6.5: The growth of E. limosum after 48 hours in PYG medium, under various gas concentrations at 70 and 80 bar. Chapter 6 196 The biomass yields obtained for 80% and 100% CO2 were much lower than those for 20% and 50% CO2. This indicates that even though E. limosum was able to grow under all the gas concentrations tested, the increase in the CO2 concentrations (with a decrease in the H2) did influence growth. Growth was also similar for both the supercritical and the subcritical conditions, indicating that the supercritical state of the CO2 did not affect the growth adversely when compared with the subcritical state. Considering Figure 6.4, the only parameters which changed is lowering the H2, while increasing the CO2, which should now reflect in the metabolic activity and thus affected the growth. Figure 6.6: The growth of E. limosum after 48 hours in various media compositions, under 100% CO2 at 70 and 80 bar. For the different media compositions, it can be seen that an increase in the biomass yields were obtained for the PYG and MM + Glc media. However, no increase in the yields were obtained when E. limosum was grown in the nutrient deprived MM – Glc medium and when only fissure water was used. This indicated that when an additional carbon source was added to the medium, E. limosum was able to grow. However, when no additional carbon source was added, E. limosum was not able to use CO2 as the sole carbon source to produce biomass when growing under >70 bar in 100% CO2. In the literature it has been described that E. limosum has the ability to grow under pressures above 1 atm (1.01325 bar) when using CO as the sole carbon or energy source (Genthner & Bryant, 1982). However, when CO2 was used as the sole carbon source, H2 was additionally added as the energy source (Genthner et al., 1981; Genthner & Bryant, Chapter 6 197 1982; Loubière & Lindley, 1991; Lelait & Grivet, 1996; Roh et al., 2011). In other experiments, glucose, methanol, or acetate was added to improve growth (Genthner et al., 1981; Loubière & Lindley). Therefore, this is the first report where it was shown that E. limosum can grow beyond 2 bar, surprisingly even up to 100 bar using CO2 as the sole carbon and energy source. E. limosum has the ability to fix CO and CO2 through the use of the Wood–Ljungdahl pathway [also known as the reductive acetyl coenzyme A (Acetyl-CoA) pathway] (Loubière & Lindley, 1991; Roh et al., 2011). According to Chivian et al., (2008), the application of the Wood–Ljungdahl pathway may have the added benefit of Na+ export to aid in maintaining the Na+ gradient utilized by the Na+ antiporters and symporters. This includes the Na+/H+ antiporter that could aid in driving ATP synthase (H+-dependent). Even though growth was low or minimal under certain conditions, the ATP measurements still displayed positive values for the cultures from all the syringe experiments. Therefore, one would expect that when E. limosum was not able to produce biomass under specific conditions, it could stay viable, but with limited metabolic activity. As additional confirmation that E. limosum stayed viable, live/dead stains (Section 3.3.6.10.2) were conducted for the different pressures (Figure 6.7), CO2 concentrations (Figure 6.8), and media compositions (Figure 6.9). 10 Bar 20 Bar 30 Bar 40 Bar Chapter 6 198 50 Bar 60 Bar 70 Bar 80 Bar 90 Bar 100 Bar Figure 6.7: Live/Dead stain images of E. limosum after 48 hours in PYG medium, under various high pressures and 20% CO2. All scale bars are equal to 2 µm. The live/dead stains images confirmed that even though some of the cells died, many E. limosum cells were still alive after being subjected to pressures up to 100 bar. 20% CO2 – 70 Bar 20% CO2 – 80 Bar Chapter 6 199 50% CO2 – 70 Bar 50% CO2 – 80 Bar 80% CO2 – 70 Bar 80% CO2 – 80 Bar 100% CO2 – 70 Bar 100% CO2 – 80 Bar Figure 6.8: Live/Dead stain images of E. limosum after 48 hours in PYG medium, under various gas concentrations at 70 and 80 bar. All scale bars are equal to 2 µm. Figure 6.8 and Figure 6.9 indicate live/dead stain activity after growth under different gas concentrations and different media compositions, respectively. PYG – 70 Bar PYG – 80 Bar Chapter 6 200 MM + Glc – 70 Bar MM + Glc – 80 Bar MM - Glc – 70 Bar MM - Glc – 80 Bar FW – 70 Bar FW – 80 Bar Figure 6.9: Live/Dead stain images of E. limosum after 48 hours in various media compositions, under 100% CO2 at 70 and 80 bar. All scale bars are equal to 2 µm. 6.4.2 Cultivation and Analyses of the Subsurface Biome Growth of the subsurface biome was analysed by optical density after sacrificing the syringe experiments incubated under different pressures (Figure 6.10), different CO2 concentrations (Figure 6.11), and different media compositions (Figure 6.12) as discussed in Section 4.3.4. Chapter 6 201 Figure 6.10: The growth of the subsurface biome after 48 hours in LB medium, under various high pressures and 20% CO2. An increase in the biomass yields was obtained for the subsurface biome after 48 hours of growth under all the different pressures tested. This indicated that even though the yields obtained were very low, the biome was able to proliferate under pressures up to 100 bar. Figure 6.11: The growth of the subsurface biome after 48 hours in LB medium, under various gas concentrations at 70 and 80 bar. The biomass yields obtained for the different gas concentrations decreased as the CO2 concentration increased. At 100% CO2, the biomass yields were almost negligible, Chapter 6 202 indicating that there was little to no growth after 48 hours. Growth was similar for both the supercritical and the subcritical conditions. As discussed for E. limosum, the lowering of the H2 is reflected in the metabolic activity and thus affected growth. Figure 6.12: The growth of the subsurface biome after 48 hours in various media compositions, under 100% CO2 at 70 and 80 bar. Studies of the subsurface biome in the different media compositions indicated that growth only occurred when the medium contained an additional carbon source. Under nutrient limited conditions, almost no increase in the biomass yield was obtained. However, even though minimal growth was observed under all these conditions, the ATP measurements were still positive for the biome from all the syringe experiments. The Wood–Ljungdahl pathway occurs in the Archaeal methanogens and acetate-producing Bacteria such as Clostridium (Matschiavelli et al., 2012). As shown in Chapter 3, both these types of microorganisms are present in the subsurface biome, therefore, one would expect that the Wood–Ljungdahl pathway may aid in driving ATP synthase. Live/dead stains (Section 3.3.6.10.2) were conducted for the different pressures (Figure 6.13), CO2 concentrations (Figure 6.14) and media compositions (Figure 6.15), to confirm that the biome was still viable. The images shown were selected to also indicate the various morphologies present in the biome. Chapter 6 203 10 Bar 20 Bar 30 Bar 40 Bar 50 Bar 60 Bar 70 Bar 80 Bar Chapter 6 204 90 Bar 100 Bar Figure 6.13: Live/Dead stain images of the subsurface biome after 48 hours in LB medium, under various high pressures and 20% CO2. All scale bars are equal to 2 µm. These images confirmed that the microorganisms were still mostly alive, even after 48 hours under pressures up to 100 bar. It was also found that as discussed in Section 3.4.3.3, the microorganisms formed biofilm-like structures through accumulation of cells, possibly as a stress response to protect against the conditions they were subjected to (Mitchell et al., 2009). 20% CO – 70 Bar 20% CO – 80 Bar 2 2 50% CO2 – 70 Bar 50% CO2 – 80 Bar Chapter 6 205 80% CO2 – 70 Bar 80% CO2 – 80 Bar 100% CO – 70 Bar 100% CO – 80 Bar 2 2 Figure 6.14: Live/Dead stain images of the subsurface biome after 48 hours in LB medium, under various gas concentrations at 70 and 80 bar. All scale bars are equal to 2 µm. Figure 6.14 and Figure 6.15 indicate live/dead stain activity after growth under different gas concentrations and different media compositions, respectively. PYG – 70 Bar PYG – 80 Bar Chapter 6 206 MM + Glc – 70 Bar MM + Glc – 80 Bar MM - Glc – 70 Bar MM - Glc – 80 Bar FW – 70 Bar FW – 80 Bar Figure 6.15: Live/Dead stain images of the subsurface biome after 48 hours in various media compositions, under 100% CO2 at 70 and 80 bar. All scale bars are equal to 2 µm. Chapter 6 207 6.5 CONCLUSIONS Supercritical CO2 generally impacts the majority of microorganisms in a negative manner (McHugh & Krukonis, 1994; Zhang et al., 2006; Mitchell et al., 2009; Basava-Reddi, 2012; Peet et al., 2015). However, in this study it was found that even though the microorganisms were not able to grow, they still remained alive under all the conditions tested, thus the subsurface biome from the Star Diamonds could successfully be evaluated in an environment that mimics geological CCS. In the literature, several Firmicutes, especially Bacillus species have been described that were resistant to sc- CO2 and were able to grow in pressurized bioreactors containing sc-CO2 (Peet et al., 2015; Peet & Thompson, 2015). It has also been reported that native microorganisms that were present at CO2 injection sites, were able to survive sc-CO2 conditions (Colwell et al., 2011). Diversity studies of the subsurface biome (Chapter 3) indicate that several Firmicutes, including Bacillus species were present in the biome, therefore, linking these observations to microorganisms from the subsurface biome that were able to tolerate and survive the sc-CO2 conditions. When studies were done on the CO2 stress response of three model organisms (Shewanella oneidensis MR-1, Geobacillus stearothermophilus, and Methanothermobacter thermoautotrophicus), it was shown that biofilm formation and cell wall thickness may be two very important factors in resisting sc-CO2 toxicity, since the presence of a biofilm can create a reactive barrier that reduces the rate of CO2 diffusion into the cytoplasmic membranes and facilitate conversion of CO2 into long-term stable carbonate phases (Gerlach et al., 2011; Santillan et al., 2011; Basava-Reddi, 2012). The live/dead images show that the biome formed biofilm-like structures through the accumulation of cells. Since the biofilm can offer structural support, protection from physical, chemical, and biological stresses, optimal location relative to substrates required for metabolic function, and symbiotic benefits in multi-species communities (Watnick & Kolter, 2000; Costerton & Stewart, 2001; Stoodley et al., 2002; Mitchell et al., 2009; Lewandowski & Beyenal, 2013), it is possible that the microorganisms were able to protect themselves from any stresses created by the sc-CO2 conditions by forming these biofilm-like structures. Additionally, biofilms can cause large changes in porosity and permeability of porous media including sandstone, but it can also act as a hydraulic Chapter 6 208 barrier to prevent the flux of CO2, making them potentially useful for preventing leakage of sc-CO2 from the injection sites (Mitchell et al., 2009). It has also been found that during nutrient starvation the vitamin, nutrient, and energy requirements of the microorganisms can no longer be fulfilled, resulting in cells that will separate themselves from the biofilm in order to find other microenvironments that have these requirements (Hunt et al., 2004; Thormann et al., 2005; Thormann et al., 2006; Mitchell et al., 2009). In order to sustain the integrity of the biofilm over time, constant feeding with sufficient nutrient and energy sources are required (Mitchell et al., 2009). When the microorganisms were subjected to the nutrient deprived media, they were not able to produce biomass, indicating that they were most likely not able to form sufficient biofilms to protect themselves against sc-CO2 and as a result they used their energy to stay alive, rather than to produce biomass. The Gram-staining properties of the microorganisms will also influence their ability to resist sc-CO2 toxicity, since Gram-positive microorganisms are more resilient towards sc- CO2 than Gram-negative microorganisms due to the high peptidoglycan content in their cell walls, making them less permeable (Cheftel, 1995; Zhang et al., 2006; Mitchell et al., 2009). In Chapter 4 it was shown that E. limosum is a Gram-positive rod (Song & Cho, 2015), and should therefore have better resistance towards sc-CO2, explaining why they were able to stay alive under the sc-CO2 conditions. The subsurface biome contained both Gram-positive and Gram-negative microorganisms (Chapter 4), therefore, microorganisms will be present that are more resilient towards sc-CO2 than others, indicating that the community will most likely change during CCS. Chapter 6 209 6.6 REFERENCES Basava-Reddi, L. (2012). Microbial Effects on CO2 Storage. IEAGHG Report. Benson, S.M. and Cole, D.R. (2008). 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Terminal Microbial Metabolisms in the Deep Subsurface under Conditions Relevant to CO2 Sequestration and Enhancing Methanogenesis from Coal. PhD Thesis, Colorado School of Mines. Chapter 6 211 Hong, S.I., Park, W.S. and Pyun, Y.R. (1997). Inactivation of Lactobacillus sp. from Kimchi by High-Pressure Carbon Dioxide. Food Science and Technology- Lebensmittel-Wissenschaft & Technologie. 30:681-685. https://en.wikipedia.org/wiki/Supercritical_carbon_dioxide. Hunt, S.M., Werner, E.M., Huang, B.C., Hamilton, M.A. and Stewart, P.S. (2004). Hypothesis for the Role of Nutrient Starvation in Biofilm Detachment. Applied and Environmental Microbiology. 70:7418-7425. Jørgensen, B.B and Boetius, A. (2007). Feast and Famine – Microbial Life in the Deep-Sea Bed. Nature Reviews Microbiology. 5:770-781. Jørgensen, B.B. and D’Hondt, S. (2006). ECOLOGY: A Starving Majority Deep Beneath the Seafloor. Science. 314:932-934. Lelait, M. and Grivet, J.P. (1996). 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Changes in the Deep Subsurface Microbial Biosphere Resulting from a Field-Scale CO2 Geosequestration Experiment. Frontiers in Microbiology. 5:209. Nondorf, L., Gutierrez, M. and Plymate, T.G. (2011). Modelling Carbon Sequestration Geochemical Reactions for a Proposed Site in Springfield, Missouri. Environmental Geosciences. 18:91-99. Onstott, T.C., Colwell, F.S., Kieft, T.L., Murdoch, L. and Phelps, T.J. (2009). New Horizons for Deep Subsurface Microbiology. Microbe Magazine. 4(11):499– 505. Chapter 6 213 Peet, K.C. and Thompson, J.R. (2015). Draft Genome Sequences of Supercritical CO2-Tolerant Bacteria Bacillus subterraneus MITOT1 and Bacillus cereus MIT0214. Genome Announcements. 3(2):e00140-15. Peet, K.C., Freedman, A.J.E., Hernandez, H.H., Britto, V., Boreham, C., Ajo- Franklin, J.B. and Thompson, J.R. (2015). Microbial Growth under Supercritical CO2. Appl Environ Microbiol. 81(8):2881-2892. Phelps, T.J., Murphy, E.M., Pfiffner, S.M. and White, D.C. (1994). Comparison between Geochemical and Biological Estimates of Subsurface Microbial Activities. Microb. Ecol. 28:335-349. Roh, H., Ko, H., Kim, D., Choi, D.G., Park, S., Kim, S., Chang, I.S. and Choi, I. (2011). Complete Genome Sequence of a Carbon Monoxide-Utilizing Acetogen, Eubacterium limosum KIST612. Journal of Bacteriology. 193(1):307-308. Rozzi, N.L. and Singh, R.K. (2002). Supercritical Fluids and the Food Industry. Comp. Rev. Food Sci. Food Safety. 1:33-44. Santillan, E.F.U., Franks, M.A., Omelon, C.R. and Bennett, P. (2011). Microbes under Pressure: A Comparison of CO2 Stress Responses on Three Model Organisms and their Implications for Geologic Carbon Sequestration. American Geophysical Union, Fall Meeting, Abstract #B51J-0552. Song, Y. and Cho, B. (2015). Draft Genome Sequence of Chemolithoautotrophic Acetogenic Butanol-Producing Eubacterium limosum ATCC 8486. Genome Announcements. 3(1):e01564-14. Stoodley, P., Sauer, K., Davies, D.G. and Costerton, J,W. (2002). Biofilms as Complex Differentiated Communities. Annual Review of Microbiology. 56:187-209. Thormann, K.M., Duttler, S., Saville, R.M., Hyodo, M., Shukla, S., Hayakawa, Y. and Spormann, A.M. (2006). Control of Formation and Cellular Detachment from Chapter 6 214 Shewanella oneidensis MR-1 Biofilms by Cyclic di-GMP. Journal of Bacteriology. 188:2681-2691. Thormann, K.M., Saville, R.M., Shukla, S. and Spormann, A.M. (2005). Induction of Rapid Detachment in Shewanella oneidensis MR-1 Biofilms. Journal of Bacteriology. 187:1014-1021. Watnick, P and Kolter, R. (2000). Biofilm, City of Microbes. Journal of Bacteriology. 182:2675-2679. White, C.M., Strazisar, B.R., Granite, E.J., Hoffman, J.S. and Pennline, H.W. (2003). Separation and Capture of CO2 from Large Stationary Sources and Sequestration in Geological Formations - Coalbeds and Deep Saline Aquifers. Journal of the Air & Waste Management Association. 53:645-715. Wilkins, M.J., Daly, R.A., Mouser, P.J., Trexler, R., Sharma, S., Cole, D.R., Wrighton, K.C., Biddle, J.F., Denis, E.H., Fredrickson, J.K., Kieft, T.L., Onstott, T.C., Peterson, L., Pfiffner, S.M., Phelps, T.J. and Schrenk, M.O. (2014). Trends and Future Challenges in Sampling the Deep Terrestrial Biosphere. Frontiers in Microbiology. 5:481. Zhang, J., Davis, T.A., Matthews M.A., Drews, M.J., LaBerge, M. and An, Y.H. (2006). Sterilization using High-Pressure Carbon Dioxide. Journal of Supercritical Fluids. 38:354-372. Chapter 6 215 CHAPTER 7 CARBON SEQUESTRATION SIMULATIONS OF THE SUBSURFACE BIOME USING A CONTINUOUS HIGH PRESSURE BIOREACTOR 7.1 INTRODUCTION Before CO2 can be stored on a scale that can make a difference in global climate change, the longevity of sequestering CO2 in the storage formation must be addressed. The temperatures and pressures required for the injection of sc-CO2 can affect the composition and metabolic activity and in turn, the metabolic cycling of a wide variety of microorganisms and could thus potentially have an impact on CO2 injection, migration, and storage longevity (Morozova et al., 2010; Glossner, 2013). Various studies on subsurface microbial communities have revealed that the more important metabolic groups, dominating the deep subsurface, are sulphate reduction, iron reduction, fermentation, and methanogenesis (Takai et al., 2001; Kotelnikova, 2002; Baker et al., 2003; Newberry et al., 2004; Onstott, 2005; Webster et al., 2006; Fry et al., 2008; Basso et al., 2009; Morozova et al., 2010; Ragon et al., 2013), thus, the presence of sc-CO2 can affect the metabolism of both heterotrophic microorganisms, which play a major role in carbon cycling, as well as the chemolithoautotrophic microorganisms which are able to use CO2 as a sole carbon and energy source (Chapelle et al., 2002; Morozova et al., 2010; Hubert et al., 2011; Bordenave et al., 2012). As discussed in Chapter 6, sc-CO2 has been shown to be harmful to several microorganisms, but has less of an effect on biofilm-forming microorganisms. Mitchell et al., (2008) and Mitchell et al., (2009), have studied the effect of sc-CO2 on biofilm microorganisms in Berea sandstone and unconsolidated sand. They found that biofilm- forming microorganisms increased the permeability of the Berea sandstone, but the sc- CO2 had a negligible effect on their biofilm formation, confirming that sc-CO2 may not completely sterilize geological environments when the microorganisms are attached to Chapter 7 216 rock surfaces. However, these studies were done with pure cultures and not much is known about the effects of sc-CO2 on mixed microbial communities, especially those found in the subsurface environments considered for CCS (Morozova et al., 2010; Glossner, 2013). Other than temperature, pressure, and the presence of sc-CO2, factors such as changes in pH and salinity, occurring due to the sequestration of CO2, may also significantly influence subsurface microbial communities (Morozova et al., 2010; Krüger et al., 2011; Santillan et al., 2013). Thus, it becomes extremely important to characterize the diversity of the deep subsurface, before, during, and after injection of CO2 since the interaction of the subsurface microbial communities within the storage rock could be responsible for significant changes in the structure and chemical composition of the rock formations and as a result, can influence the porosity and permeability of the storage reservoir and leakage of the CO2 (Morozova et al., 2010; Krüger et al., 2011; Santillan et al., 2013). This leakage can affect the quality of the ground water through acidification of the water, resulting in the mobilization of metals to toxic concentrations, which can influence the native microbial communities (Krüger et al., 2011). 7.2 AIMS OF THIS CHAPTER The main aims of this chapter were to: Evaluate the potential impact of CO2 sequestration on the subsurface microbial communities through bench-scale simulations of these interactions using a continuous high pressure bioreactor. Determine the impact on the diversity within the selected biome and define possible CO2 fixation pathways that the subsurface biome might use during CCS interaction. Chapter 7 217 7.3 MATERIALS AND METHODS For all the high pressure experiments in this phase, the bioreactor described in the final design (Section 5.3.2), was used. The bioreactor was sterilized and prepared as discussed in Section 5.3.2.2. Unless otherwise specified, the fissure water containing the subsurface biome was from the Nalgene Carboy bottle that contained large amounts of cells after collection on the massive filter as discussed in Section 3.3.6.4. Crushed sandstone, prepared as discussed in Section 3.3.5, was used as the rock matrix inside the bioreactor. 7.3.1 Adhesion Studies Adhesion studies were carried out to determine the amount of time needed for the microorganisms from the subsurface biome to adhere to the crushed rock before the reactor was circulated. This was done by filling 20 ml syringes with the crushed sandstone. The syringes were wrapped in foil and sterilized. After sterilization, the syringes were filled with fissure water containing the subsurface biome and covered at both ends with parafilm (Lasec) (Figure 7.1). Figure 7.1: The prepared syringes, used for the adhesion studies. At each specified time interval, triplicate syringes were sacrificed by draining the syringe and collecting the liquid in a sterile 15 ml falcon tube. To determine if the microorganisms adhered to the rock, live/dead stains using the LIVE/DEAD® BacLightTM Chapter 7 218 Bacterial Viability Kit (Molecular Probes, Inc., Invitrogen, Promega) (Section 3.3.6.8.2), and ATP measurements using the CellTiter-Glo® Luminescent Cell Viability Assay (Promega) were done according to the manufacturer’s instructions. This was repeated at various time intervals, until no cells were released and the corresponding ATP values were negative, ensuring complete adhesion of the microorganisms. The time of complete adhesion was determined to be 24 hours, but the adhesion studies were continued for 48 hours, to confirm that the microorganisms remained attached to the sandstone particles. 7.3.2 Carbon Sequestration Simulations with the Subsurface Biome The sterilized bioreactor was packed with the crushed sandstone, where after the fissure water, containing the subsurface biome was added to the bioreactor until the entire volume was filled as discussed in Section 5.3.2.2. This was left to stand for 24 hours, as determined by the adhesion studies (Section 7.3.1), in order for the microorganisms to adhere to the rock. CO2 gas was then added to saturate the system and to achieve the initial pressure (~50 bar). The final pressure of ≥80 bar, to allow the CO2 to be in a supercritical state, was achieved and maintained by pressurizing the piston pressure regulators with the nitrogen gas and the valves were opened to allow circulation as discussed in Section 5.3.2.2. No additional carbon and energy sources were added, to simulate the subsurface CO2 storage conditions. The temperature was maintained at 35°C and the reactor was allowed to run for six weeks. During the six weeks, 1 ml samples were taken at various time intervals for continual analyses to determine if any metabolic cycling occurred inside the bioreactor under CCS conditions. As discussed in Section 5.4, the bioreactor design only allowed for a total system volume of ~400 ml, therefore, only limited samples could be withdrawn and thus analyses were restricted. Sampling was done with a gas tight syringe as discussed in Section 5.3.2.2. Half of the liquid sample (500 µl), as well as any gas that might have formed inside the syringe headspace was immediately added to a sterile 15 ml serum vial, which was sealed with a blue chlorobutyl stopper (Bellco Glass Inc.) and crimp sealed with an aluminium cap. This was used to test the gas composition, especially the CO2 usage using a GC-2010 Plus capillary gas chromatograph (GC), coupled to a Chapter 7 219 Barrier Discharge Ionization (BID) detector (Shimadzu). A micropacked ShinCarbon ST 80/100 column (Restek®), with an inner diameter of 0.53 mm and 2 m in length, was used. Gas samples were introduced into the system by means of a manual injection valve fitted with a 20 µL loop. The injector port temperature was 80°C in the splitless mode and the carrier gas was helium (99.9999%). The initial oven temperature was 40°C held for 2 minutes and then raised at 20°C/minute to 140°C. Detection was carried out with a BID detector set a 280°C and with a plasma gas flow of 80 ml.min-1. Instrument control, data collection, and data processing were carried out with Shimadzu LabSolutions software (V.5.57). For the other 500 µl of the sample, 100 µl was inoculated into a sterile Hungate tube containing 5 ml LB medium (Section 4.3.4) and incubated at 35°C, as a visual test, to determine of the microorganisms inside the bioreactor were still viable. The remaining 400 µl was first used to measure the pH, where after it was sonicated for 10 minutes, to allow breakage of the cell walls of the microorganisms in order to detect any intracellular product formation. The cell suspension was centrifuged (20 000 x g, 10 minutes, 4°C) using an AllegraTM 25R centrifuge (Beckman CoulterTM) and the supernatant was added to a high performance liquid chromatograph (HPLC) vial to be used for the detection of any saccharide or acid formation. HPLC analyses were carried out on a system consisting of an isocratic pump, an auto sampler, and a photodiode array detector (Finnegan Surveyor). Analytes were separated on an Aminex HPX-87H (7.8 x 300 mm) ion exchange column (Bio-Rad Laboratories, Inc.), maintained at 65°C. Elution was performed with 5 mM sulphuric acid at a flow rate of 0.5 ml.min-1. Organic acids were detected at 202 and 276 nm and saccharides were detected with a 2414 Refractive Index detector (RID) (Waters) connected in series. RID cell temperature was 40°C and data recording and analyses were carried out with the Clarity software package (DataApex). Analyses were also done on samples that were taken from the fissure water before it was inserted into the bioreactor, from the bioreactor after adhesion occurred, but before the addition of the CO2 gas, as well as immediately after the addition of the CO2 gas. After the six weeks, circulation of the bioreactor was stopped and microbial diversity analyses using DGGE (Section 3.3.6.6), as well as geochemical analyses were conducted on the remaining liquid. Cations were measured using an ICP Optima 3000 DV (PerkinElmer Inc.), and anions were analysed using an IC DX-120 (Dionex). Chapter 7 220 Geochemical analyses were done at the Institute for Ground Water Studies (IGS) at the UFS (Bloemfontein, RSA). The composition of the sandstone was analysed with XRF (Section 3.3.5.2) at the Department of Geology at the UFS (Bloemfontein, RSA). The microbial diversity in the bioreactor was visualized using SEM, with EDS analyses (Section 3.3.6.8.4) at the Centre for Microscopy at the UFS (Bloemfontein, RSA). The bioreactor study was repeated using the same conditions as before, except the CO2 13 13 gas used to saturate the system was labelled with the C isotope ( CO2) and consisted of 99 atom % 13C, with <3 atom % 18O (Sigma-Aldrich®). This was done to detect possible incorporation of the 13C in the DNA using Nuclear Magnetic Resonance (NMR) to indicate metabolic fixation of the CO2 by the microorganisms. Unfortunately, this technique required high biomass concentrations and since this could not be obtained under the CCS conditions and current bioreactor conditions, as indicated in Chapters 4 and 6, it was not possible to carry out analyses using NMR. In the subsurface, the microorganisms are constantly in contact with a fresh fissure water supply, thus, in order to better simulate the subsurface CCS conditions, additional fissure water was added to the reactor every second day, to ensure that any nutrients from the fissure water did not become depleted. After six weeks, microbial diversity assessments using targeted rRNA gene sequencing, was done on the Ion Torrent PGMTM as discussed in Section 3.3.6.7. Sequencing was done at the CAF, Stellenbosch University (Stellenbosch, RSA). 7.3.3 Bioreactor Control Experiments As discussed in Chapter 5, the material used in constructing the bioreactor was made out of corrosion-free stainless steel, but any potential corrosion and metal leakage from the bioreactor was considered. Thus; to determine the degree to which metals would leach from the reactor itself, if at all, several negative control experiments were done. All these experiments were carried out in the absence of the subsurface biome. Control experiments continued for two weeks under specific conditions, as indicated in Table 7.1. Chapter 7 221 Table 7.1: Conditions for the bioreactor control experiments. Component A1 A2 B1 B2 B3 Fissure water + + + + + Sandstone - - + + + CO2 - + - - + Pressure (Bar) 0 80 0 80 80 Legend: + = component was added to the reactor, - = component was not added to the reactor. After the two weeks, the water chemistry and sandstone composition was analysed for each control as discussed in Section 7.3.2. These analyses were compared with fissure water and sandstone that was not in contact with the bioreactor. 7.3.4 Whole Transcriptome Sequencing using Ion Torrent ProtonTM The diversity studies in Chapter 3 indicated that carbon fixing microorganisms are present in the subsurface biome. Thus, in order to define possible CO2 fixation pathways that these microorganisms might follow if they survive the CCS conditions, whole transcriptome sequencing was done on the subsurface biome. 7.3.4.1 Total RNA Isolation Total ribonucleic acid (RNA) was isolated from the massive filters (Section 3.3.6.4), the PVC cartridges (Section 3.3.6.5), and the second bioreactor (Section 7.3.2) using two different methods. The first method was according to the procedures of Lau et al., (2014) as discussed in Section 3.3.6.7.1, but by adding rDNase (Macherey-Nagel) to the re- suspended pellets. The second method used was with the NucleoSpin® RNA kit (Macherey-Nagel) according to the manufacturer’s instructions. The primers and standards used were as recommended by the manufacturer. 7.3.4.2 RNA Sequencing and Analyses The RNA was used for RNA sequence analyses using the Ion Torrent ProtonTM platform (Life TechnologiesTM), by constructing a whole transcriptome library from the RNA for up Chapter 7 222 to 200-base-read sequencing according to the manufacturer’s instructions. The main steps in constructing the library included hybridizing and ligating the RNA, reverse transcription (RT) to make complementary DNA (cDNA), purification and amplification of the cDNA, purification of the amplified cDNA, and assessment of the yield and size distribution of the amplified cDNA. The template was prepared from the whole transcriptome library and sequenced using the Ion PITM Hi-QTM OT2 200 Kit (Thermo Fisher Scientific) according to the manufacturer’s instructions. Sequencing reactions were done with the Ion Torrent ProtonTM (Life TechnologiesTM) at the CAF, Stellenbosch University (Stellenbosch, RSA). The data was analysed with MG-RAST using the default settings (Meyer et al., 2008). Chapter 7 223 7.4 RESULTS AND DISCUSSIONS 7.4.1 Carbon Sequestration Simulations with the Subsurface Biome Immediately after the addition of the CO2 gas, a drop in the pH was observed (Table 7.2) due to the formation of carbonic acid as the CO2 dissolves in the water (Fierer & Jackson, 2006; Mu et al., 2014). The presence of sc-CO2 or carbonic acid can lead to reactions with the minerals in the storage formations, resulting in dissolved ions which could in turn lead to mineral dissolution or precipitation. This could influence the ionic strength and the distribution of bioavailable terminal electron acceptors such as Fe(III), affecting microbial metabolism in the subsurface (McMahon & Chapelle, 1991; Cozzarelli et al., 1994; Banfield et al., 1999; Rogers & Bennett, 2004), but also the geochemistry and mineralogy of the storage rock, changing its porosity, and leading to possible acetogenesis and methanogenesis (Ragsdale, 2004; Gadd, 2010; Sato et al., 2013). However, other than this initial drop, no dramatic changes in the pH were observed during the six weeks. This potentially indicated that products such as acetate, which would have caused a further decrease in the pH, or carbonates, which would have caused an increase in the pH, were not formed in amounts significant enough to cause a shift in the pH. Table 7.2: pH and growth measurements obtained of the bioreactor during the six weeks. Sample name Date pH Growth obtained Fissure water 27/11/2014 7.44 + Before the addition of CO2 28/11/2014 7.29 + After the addition of CO2 28/11/2014 5.43 + S1 02/12/2014 5.48 + S2 05/12/2014 5.53 + S3 09/12/2014 5.63 + S4 17/12/2014 5.63 + S5 20/12/2014 5.56 + S6 24/12/2014 5.57 + S7 29/12/2014 5.58 + S8 03/01/2015 5.61 + Legend: + = growth was obtained for the sample. The pH measurements for the second bioreactor followed the same trends from 23/01/2015 until 05/03/2015. However, the pH was not measured under pressure and is Chapter 7 224 therefore just a possible indication of a trend that was followed, not an actual statement of the supercritical conditions inside the bioreactor. It is known that changes in environmental parameters such as a decrease in the pH, could affect the microorganisms and their cellular and biochemical functions, influencing their activity and composition (Booth et al., 2002; Morozova et al., 2010; Basava-Reddi, 2012), but it can also allow growth, and thus dominance of acid tolerant species (Fierer & Jackson, 2006). Therefore, each time the pH was measured, the sample was also inoculated for growth as discussed in Section 7.3.2. It was found that even though no further changes in the pH were observed, the microorganisms remained viable, even after six weeks subjected to CCS conditions, since they were able to grow when revived in the LB medium. But the effect which the CO2 has on pH could influence the microbial communities present in the subsurface biomes. Since the diversity studies in Chapter 3 revealed that both the methanogens and the acetogens such as Clostridium, are present within the subsurface biome, this might potentially lead to methanogenesis, however, according to Basava-Reddi, (2012), the extent to which the decreased pH will affect the microbial communities, depends on the microorganisms present, the geology and mineralogy of the storage rock, as well as the chemistry of the water. Thus, product formation such as acetate and methane, could be an indication of potential methanogenesis. The GC results (Figure 7.2) indicated that after six weeks, no major differences were observed in the compositional gas analyses and that the formation of methane was not detected. It is however possible that since the reactor was over saturated with CO2, the GC analyses were not sensitive enough to detect changes, indicating minor usage thereof. Since results were constant during the six weeks, analyses for the rest of the samples are not shown. As discussed in Section 7.3.2, the sample used for the GC analyses was immediately added to a sterile 15 ml serum vial. This vial was not evacuated before injection; therefore, prominent N2 and O2 peaks were detected in the analyses. The HPLC results also did not yield any products after six weeks subjected to CCS conditions, with no real differences observed for the formation of acetate and formate, also no saccharides such as glucose were detected. Chapter 7 225 N2 O2 CO2 H2 CO Methane Ethane A B C Figure 7.2: GC analyses of (A) fissure water, before the addition of CO2, (B) after the addition of CO2, and (C) S8 - after 6 weeks. 7.4.2 Geochemical Analyses Geochemical analyses of the bioreactor water were conducted as discussed in Section 7.3.2, and the data is displayed in Table 7.3. When the geochemical results from the bioreactor water were compared with the geochemical analyses of the fissure water (Section 3.4.1.2), it can be seen that in general the composition remained relatively constant, indicating that the bioreactor water and the fissure water stayed geochemically very similar (Figure 7.3), except for the increase in aluminium, iron, and M-alkalinity in the bioreactor water after six weeks (Figure 7.4). Chapter 7 226 Table 7.3: Geochemical analyses of the bioreactor. Analyses: Bioreactor pH 5.6 EC (mS.m-1) 461 TDS (mg.L-1) 2832 P-Alkalinity (mg.L-1) 0 M-Alkalinity (mg.L-1 425 Calcium Hardness (mg.L-1) 239 Magnesium Hardness (mg.L-1) 39 Total Hardness (mg.L-1) 279 Aluminium as Al (mg.L-1) 2731.5 Barium as Ba (mg.L-1) 0.516 Bromide as Br (mg.L-1) 7.30 Cadmium as Cd (mg.L-1) <0.002 Calcium as Ca (mg.L-1) 95.7 Chloride as Cl (mg.L-1) 1428 Cobalt as Co (mg.L-1) 0.213 Copper as Cu (mg.L-1) <0.002 Fluoride as F (mg.L-1) 0.86 Iron as Fe (mg.L-1) 97.99 Lead as Pb (mg.L-1) 0.012 Magnesium as Mg (mg.L-1) 9.6 Manganese as Mn (mg.L-1) 1.51 Potassium as K (mg.L-1) 11 Sodium as Na (mg.L-1) 743.5 -1 Sulphate as SO4 (mg.L ) 14.12 Zinc as Zn (mg.L-1) 2.90 Legend: EC = electrical conductivity, TDS = total dissolved solids, P-Alkalinity = phenolphthalein alkalinity, M-Alkalinity = total alkalinity. The P-alkalinity was zero, thus the M-alkalinity represents bicarbonate, indicating possible formation of bicarbonate inside the bioreactor. Microorganisms can execute carboxylation and decarboxylation reactions, interconverting inorganic and organic carbon, and is extremely important in the biogeochemical cycle of carbon, since each of these reactions are responsible for a global flux of at least 10 Pmol of carbon per year (Raven, 2006; Ramanan et al., 2009). Depending on the enzyme involved, the − carboxylases use either CO2 or HCO3 . Carbonate precipitation can be achieved by an - enzymatic reaction with both CO2 and HCO3 . This enzyme, known as carbonic anhydrase (CA) is a zinc metalloenzyme, found in basically every animal and plant and - in certain bacteria and catalyses the conversion of CO2 to HCO3 (hydration) and the reverse reaction (bicarbonate dehydration). CA is amongst the fastest catalytic enzymes known (turnover values close to one million bicarbonate molecules per second) and is usually stable, even up to 85°C (Zaihua, 2001; Lane et al., 2005; Raven, 2006; Ramanan et al., 2009). Chapter 7 227 Figure 7.3: A Durov diagram of the geochemical analyses of the fissure water and the bioreactor water after six weeks. Figure 7.4: Comparisons of the geochemical analyses for the bioreactor with the fissure water. The bioreactor displayed an increase in the aluminium, the iron and the M-alkalinity. Chapter 7 228 7.4.3 X-Ray Fluorescence Analyses Major-element XRF analyses (Section 3.3.5.2) of the sandstone from the bioreactor was conducted after six weeks and compared with the XRF analyses of the sandstone before it was inserted into the bioreactor (Section 3.4.2). Results of this comparison are shown in Table 7.4. Table 7.4: Major-element XRF analyses of the sandstone before and after six weeks inside the bioreactor. Bioreactor Analyses: Sandstone Sandstone Al2O3 (%) 20.81 18.76 Fe2O3 (%) 1.065 0.89 CaO (%) 0.064 0.15 MgO (%) 0.17 0.49 MnO (%) 0.012 0.01 Na2O (%) 0 0.45 K2O (%) 2.64 2.63 P2O5 (%) 0.039 0.04 TiO2 (%) 1.103 1.01 SiO2 (%) 74.01 70.15 LOI (%) 0.087 5.42 Total (%) 100 100 Small increases were observed in the bioreactor sandstone for sodium, magnesium, iron, and calcium. But the most important difference obtained was the decrease in the aluminium after six weeks in the bioreactor, indicating that the increase in the aluminium detected in the bioreactor water with the geochemical analyses was due to the aluminium leaching from the sandstone under CCS conditions, most probably from kaolinite. Microorganisms are constantly interacting with inorganic ions. These ions can be important for their biological functions or can have inhibitory effects on their growth such as aluminium - the most abundant metal in the Earth's crust in the form of oxides and silicates (Piña & Cervantes, 1996). It has been shown in the literature that the presence of aluminium can result in toxic effects on several microorganisms (Driscoll, 1985; Meyer et al., 1994; Hervieux et al., 2002; Mecteau et al., 2002; Amonette et al., 2003; Illmer & Erlebach, 2003), usually through binding to the cell wall, causing a decrease in the permeability (Bradley & Parker, 1968; Johnson, 1988). Other mechanisms include: 1.) replacing divalent metal complexes such as magnesium and calcium in the cells and Chapter 7 229 their membranes (Amonette et al., 2003), 2.) inactivating enzymes (Illmer & Erlebach, 2003), 3.) osmoregulative disorder (Illmer & Erlebach, 2003), 4.) altering lipid-protein interactions and modifying transport activity (Suhayda & Haug, 1986; Zambenedetti et al., 1994), or 5.) complexing with ATP, DNA, and phosphates, resulting in a deficiency of the phosphate (Haug, 1984; Pettersson et al., 1985; Wood, 1995). Yaganza et al., (2004), showed that bacterial mortality rates were high, with damaged cell membranes, after exposure to 0.2 M aluminium, for 10 minutes, but at concentrations of 0.1 M and 0.05 M, membrane damage did not occur. Guida et al., (1991), found that growth of E. coli in the presence of aluminium was inhibited at 0.9 mM (pH 5.4) and at 2.25 mM (pH 6.6) and the toxicity of the aluminium increased when iron was removed from the medium, suggesting that the uptake of aluminium involves the iron transport systems (Davis et al., 1971). Thus, the toxic effects which aluminium can have on microorganisms, depends on the pH, the concentration of the aluminium, the time of exposure, as well as the presence of other metals. Aluminium usually adsorbs to mineral surfaces and when the pH is close to neutral, hydroxide complexes are formed which are insoluble, making the bioavailability of aluminium in soils and its concentrations in water very low (Piña & Cervantes, 1996). However, a decrease in the pH of soil and water, through acid rain, incorrect agricultural procedures, or disposal of industrial, mining, and domestic wastes, can result in the aluminium being solubilized and released from its natural reservoirs (Foy et al., 1978; Keyser & Munns, 1979; Ganrot, 1986; Martin, 1986; Macdonald & Martin, 1988; Myrold & Nason, 1992; Gagen et al., 1993; Stenson et al., 1993; Rogers & Bennett, 2004). After six weeks inside the bioreactor, the aluminium concentration increased to 0.1 M, at a pH of 5.6. This increase in the aluminium concentration was most likely due to the acidic and pressurized environment created by the CCS conditions, which allowed the metals to leach from the sandstone (Piña & Cervantes, 1996). Therefore, the effect of the subsurface microorganisms should be characterized. The subsurface biome was grown in LB medium containing various aluminium concentrations (0.005 M – 0.5 M) at both pH 7.0 and 5.6, and the OD was measured over time as discussed in Section 4.3.3. The pH did not influence the tolerance to aluminium and 0.2 M was the maximum aluminium concentration allowing growth. Higher concentrations resulted in detrimental effects on the subsurface biome. The µmax for each concentration was calculated as Chapter 7 230 discussed in Section 4.3 and it was observed that an increase in the aluminium concentration showed a marked decrease in the µmax. The control experiment without -1 aluminium, showed a µmax of 0.152 h . As representative values, the 0.08 M aluminium -1 -1 concentration test also resulted in a µmax of 0.152 h , the 0.1 M had a µmax of 0.120 h , -1 -1 the 0.2 M had a µmax of 0.066 h , and the 0.5 M had a µmax of 0.011 h . Therefore, the 0.1 M aluminium which the biome will encounter in the bioreactor, will slow its growth rate significantly. To determine the true effect that the aluminium might have under CCS conditions on the subsurface biome over time, the aluminium concentrations should be followed continuously inside the bioreactor during the experiments. 7.4.4 Bioreactor Control Experiments To confirm that the soluble aluminium and iron detected in the water from the bioreactor leached from the rock under acidic and higher pressure conditions, and not from the reactor material, negative control experiments were done as discussed in Section 7.3.3. Results for the geochemical analyses of the water are shown in Table 7.5. Parameters with values always below the detection limits are not shown. Table 7.5: Geochemical analyses for the fissure water and the control experiments. Analyses: FW A1 A2 B1 B2 B3 pH 7.38 7.31 5.67 7.36 7.31 5.63 EC (mS/m) 444 450 424 450 424 429 TDS (mg/L) 2210 2220 2210 2230 2200 2210 Aluminium as Al (mg/L) 0.06 0.08 0.07 0.21 1.98 1.4 Boron as B (mg/L) 2.85 2.89 2.6 2.93 3.08 3.06 Barium as Ba (mg/L) 0.77 0.78 0.73 0.32 0.27 0.24 Calcium as Ca (mg/L) 56.6 56.6 53.2 49.2 55.4 51.7 Iron as Fe (mg/L) <0.05 <0.05 <0.05 0.05 10.4 0.28 Potassium as K (mg/L) 8.6 8.9 7.8 11.4 12.1 12.0 Lithium as Li (mg/L) 0.73 0.8 0.7 0.98 1.08 1.09 Magnesium as Mg (mg/L) 1.40 1.40 1.4 5.20 6.70 5.9 Manganese as Mn (mg/L) <0.05 <0.05 0.11 0.25 0.66 0.25 Sodium as Na (mg/L) 715.5 749.0 715.3 704.9 770.3 802.6 Nickel as Ni (mg/L) <0.05 <0.05 <0.05 0.31 1.83 0.36 Silicon as Si (mg/L) 8.15 8.29 7.41 8.19 14.01 10.64 Strontium as Sr (mg/L) 3.55 3.54 3.33 2.94 3.26 3.08 Zinc as Zn (mg/L) <0.05 <0.05 0.07 0.38 2.20 0.2 Chloride as Cl (mg/L) 1856.4 1781.8 1754.1 1809.6 1428.5 1881.4 Legend: FW = fissure water, EC = electrical conductivity, TDS = total dissolved solids. When the controls were compared with the FW, a drop in pH was again observed, but only for the controls containing CO2 (A2 and B3), under 80 bar pressure. In A1, no Chapter 7 231 noticeable changes were observed, thus the bioreactor itself did not contribute to the increase in the metal concentrations, even though pressure was applied (A2), no increase in the metals was observed. The B-series controls that contained sandstone, especially when pressure was added (B2 and B3), demonstrated that metals can be released from the sandstone. The XRF results of the sandstone (Table 7.6), confirmed that the aluminium and iron leached from the sandstone. This should be considered during carbon sequestration, and especially the influence it will have on the composition and cycling of the subsurface microbial communities. Table 7.6: XRF analyses for the sandstone and the control experiments. Analyses: Sandstone B1 B2 B3 Al2O3 (%) 20.81 18.2 18.2 18.4 Fe2O3 (%) 1.065 0.9 0.9 0.8 CaO (%) 0.064 0.1 0.1 0.1 MgO (%) 0.17 0.4 0.5 0.4 MnO (%) 0.012 0 0 0 Na2O (%) 0 1.8 1.9 1.8 K2O (%) 2.64 2.6 2.6 2.6 P2O5 (%) 0.039 0 0 0 TiO2 (%) 1.103 0.8 0.8 0.7 SiO2 (%) 74.01 71.0 70.6 70.0 LOI (%) 0.087 4.2 4.3 5.0 Total (%) 100 100 100 100 7.4.5 Microbial Diversity Assessments using Denaturing Gradient Gel Electrophoresis 7.4.5.1 Genomic DNA Isolation and Polymerase Chain Reaction Genomic DNA was isolated from the contents of the second bioreactor after six weeks (Section 3.3.6.6.1) and used to amplify the Archaeal 16S (~600 bp) (Figure 7.5B), Bacterial 16S (~1500 bp) (Figure 7.5C), and Eukaryal 18S (~1700 bp) (Figure 7.5D) rRNA gene fragments as discussed in Section 3.3.6.6.2. Reactions were done in triplicate. Chapter 7 232 1 2 3 4 5 6 A B 1 2 3 4 5 6 1 2 3 4 5 6 C D Figure 7.5: (A) The O’GeneRulerTM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons, with the amplified rRNA gene fragments for (B) Archaea, (C) Bacteria, and (D) Eukarya. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. Lanes 4 – 6 contain the amplicons from the bioreactor. Amplicons were obtained for the Archaeal and Bacterial domains, but not for the Eukarya, confirming the statement made in Section 4.4.5.3, that the Eukarya will most likely not be able to survive and adapt to CCS conditions and therefore will probably not play a major role, if any, in carbon cycling in the subsurface. 7.4.5.2 Nested Polymerase Chain Reaction The purified PCR products were used to amplify the V3/V4 hypervariable regions for Archaea (~600 bp) (Figure 7.6B) and Bacteria (~600 bp) (Figure 7.6C) using a nested Chapter 7 233 DGGE PCR as discussed in Section 3.3.6.6.3. Amplicons were obtained for the V3/V4 hypervariable regions from all the samples. 1 2 3 4 5 6 1 2 3 4 5 6 A B C Figure 7.6: (A) The O’GeneRuler TM DNA Ladder Mix (Thermo Scientific) used to determine the size of the amplicons with the amplified V3/V4 hypervariable regions for (B) Archaea and (C) Bacteria. In lane 1, the O’GeneRulerTM DNA Ladder Mix is shown. Lanes 2 and 3 contain the positive and negative controls, respectively. Lanes 4 – 6 contain the amplicons from the bioreactor. 7.4.5.3 Denaturing Gradient Gel Electrophoresis, Sanger Sequencing and Analyses The DGGE PCR products were used to construct a preliminary diversity profile for Archaea (Figure 7.7A) and Bacteria (Figure 7.7B) as discussed in Section 3.3.6.6.4. 1 2 3 1 2 3 A B Figure 7.7: DGGE diversity profiles of (A) Archaea and (B) Bacteria. Chapter 7 234 The DGGE profiles obtained for the Archaea and the Bacteria after six weeks under CCS conditions inside the bioreactor, was different than what was initially obtained for the subsurface biome (Section 3.4.3.1.3), probably confirming that carbon sequestration will influence the diversity of the subsurface biome. Sequencing and analyses were conducted on selected DGGE bands as discussed in Sections 3.3.6.6.5 and 3.3.6.6.6. The analysed sequences indicated that the Bacteria were still dominated by the Proteobacteria phylum, mainly consisting of the Beta- and Gammaproteobacteria classes. This was followed by the Firmicutes phylum, consisting of the Clostridia class, the Bacteriodetes phylum, containing the Actinobacteria, Flavobacteria, and Sphingobacteriia classes, and the Deinococci class of the Deinococcus-Thermus phylum which is highly resistant to environmental hazards (Brooks & Murray, 1981; Albuquerque et al., 2005; Ekman et al., 2011). The Archaea only contained the Euryarchaeota phylum, consisting of the Methanobacteria and the Halobacteria classes. This confirmed that an overall decrease in the diversity was seen after exposure to CCS conditions. 7.4.6 Microbial Diversity Assessments using Targeted rRNA Gene Sequencing Targeted rRNA gene sequencing was done using the Ion Torrent PGMTM as discussed in Section 3.3.6.7. Extending the DGGE results, it can be seen that even though a substantial decrease in the overall diversity was observed (Figure 7.8) the Proteobacteria, especially the Betaproteobacteria, remained the dominant group, but the Firmicutes and the Bacteroidetes were also present (Figure 7.9). In Chapter 6 it was discussed that Gram-positive microorganisms are more resilient towards sc-CO2 than Gram-negative microorganisms due to the high peptidoglycan content in their cell walls, making them less permeable (Cheftel, 1995; Zhang et al., 2006; Mitchell et al., 2009). However, most of the orders present in the bioreactor were from the Gram-negative group, most probably indicating the importance of biofilm formation of the subsurface microorganisms in order to protect themselves from the stressful conditions they will be subjected to during carbon sequestration. Chapter 7 235 A (derived from Cyanobacteria) (derived from unclassified sequences) B C Figure 7.8: MG-RAST taxonomic hits distribution of the diversity in the subsurface biome for (A) class and (B) order, with (C) a rarefaction curve indicating the annotated species richness. Chapter 7 236 Figure 7.9: MG-RAST Krona diagram of the Bacteria at order level. Chapter 7 237 7.4.7 Microbial Diversity Assessments using Visual Methods 7.4.7.1 Scanning Electron Microscopy Scanning electron microscopy was conducted as discussed in Section 3.3.6.8.4. Images were falsely coloured using Adobe® Photoshop (V.CS5) and EDS analyses confirmed that the biofilm was indeed biological material. These images suggest that even though there was a substantial decrease in diversity, the biome was still composed of diverse microorganisms with varying morphology and distinctive structures (Figure 7.10). Figure 7.10: Scanning electron microscopy images of the biome after being subjected to CCS conditions. Scale bars are equal 1 µm. However, several tube-like structures (Figure 7.11) were now found to be present which were not prominent in the subsurface biome from the Star Diamonds mine. The formation of these structures could possibly be biofilm formation which takes place under Chapter 7 238 high pressures. Also, more hyphae-like structures were observed, however, the hyphae appeared to be damaged, likely due to the presence of sc-CO2, the metals, or high pressure. This suggests that, even though the microorganisms can protect themselves from these conditions to an extent, it may have detrimental effects on the subsurface biome. Figure 7.11: Scanning electron microscopy images, indicating the tube-like and hyphae-like structures. Scale bars are equal to either 1 µm or 10 µm. Chapter 7 239 EDS analyses also detected the presence of aluminium precipitation (Figure 7.12), most likely from the interaction with metals leached from the sandstone. Spectrum 40 is shown as a representative sample. Figure 7.12: EDS analyses, confirming the presence of aluminium precipitation. Chapter 7 240 7.4.8 Whole Transcriptome Sequencing using Ion Torrent ProtonTM Total RNA was isolated from the massive filters, the PVC cartridges, and the second bioreactor for whole transcriptome sequencing as discussed in Section 7.3.4. Unfortunately, not enough biomass could be obtained under the bioreactor conditions to allow proper RNA isolation. The RNA that was obtained from the bioreactor was of poor quality and could not be used for sequencing. From the diversity studies on the subsurface biome (Chapter 3), it was found that microorganisms with various metabolisms were present (Section 3.4.3.2), and included the methanogens that can fix CO2, several sulphate-reducing bacteria, including the Desulfobacterales, Desulfovibrionales, and Syntrophobacterales orders, microorganisms known to fix nitrogen such as the Cyanobacteria and Chlorobi phyla, as well as the Pseudomonadales, Rhizobiales, and Actinomycetales orders. Possible pathways that these microorganisms might follow during carbon sequestration can be divided into two general groups, as described in Bar-Even et al., (2012). The first group contains pathways such as the reductive acetyl-CoA pathway, also known as the Wood–Ljungdahl pathway, which directly utilize reduced C1 compounds (Figure 7.14) (Bar-Even et al., 2012). With the reductive acetyl-CoA pathway, the CO2 is reduced to produce CO using CO dehydrogenase. The CO combines with a methyl group to produce acetyl-CoA, via acetyl-CoA synthase (Figure 7.13) (Ljungdahl & Wood, 1965; Ragsdale, 2006; Lindahl, 2009; Ljungdahl, 2009) and occurs in acetate-producing Bacteria such as Clostridium, and the Archaea such as the methanogens (Matschiavelli et al., 2012). Thus, this may lead to the production of acetate and finally methane if hydrogen is available. Both of these groups have been found to be present in the subsurface biome, as well as after the bioreactor conditions. Chapter 7 241 Figure 7.13: The C1 carbon fixation pathways cycles [Taken from Bar-Even et al., (2012)]. The second group contains pathways such as the reductive TCA cycle, also known as the reverse Krebs cycle, through which CO2 is integrated into the carbon backbone of other metabolites, thus, reduced C1 compounds are not part of the pathway (Figure 7.13) (Bar-Even et al., 2012). Chapter 7 242 Figure 7.14: The acetyl-CoA-succinyl-CoA carbon fixation cycles [Taken from Bar-Even et al., (2012)]. The Calvin cycle, which is responsible for the fixation of most of the carbon in the biosphere, can be used to fix CO2 in order to produce glyceraldehyde-3-phosphate (G3P) which can be used to synthesize hexose sugars such as glucose, the main nutrient source for heterotrophic microorganisms (Berg et al., 2002). Chapter 7 243 Results from the whole transcriptome sequencing of the subsurface biome confirmed that sulphur metabolism, nitrogen metabolism, and CO2 fixation are present (Figure 7.15). For the CO2 fixation metabolism (Figure 7.16), it was found that gene products from the Calvin cycle were present, in particular, ribulose-1,5-biphosphate carboxylase/oxygenase (RuBisCO). RuBisCO is the predominant enzyme in carbon fixation and the rate limiting enzyme in the Calvin Cycle and is a carbonic anhydrase (Feller et al., 2008; Yeates et al., 2008), therefore, the production of bicarbonates are possible (Zaihua, 2001; Lane et al., 2005; Raven, 2006; Ramanan et al., 2009). The carboxysomes also contain the RuBisCO enzyme. Some of the more important enzymes involved in the C1 carbon fixation pathways present in the subsurface biome, included formate dehydrogenase, CO dehydrogenase, and acetyl-CoA synthase. The acetogens, present within the subsurface biome, as well as after the bioreactor conditions, can reduce CO2 to formate using formate dehydrogenase (Yamamoto et al., 1983; Ljungdahl, 1986). The acetogens can also synthesize CO from CO2 using ferredoxin as the electron donor and they can use acetyl-CoA synthase to generate acetyl-CoA (Drake et al., 2006; Ragsdale, 2008). Other gene products involved in central carbohydrate metabolism such as the gene products for hydroxyacylglutathione hydrolase, were also present. Thus, one might anticipate that the subsurface biome should be able to produce formate, acetate, methane, and bicarbonates under CCS conditions. The diversity studies on the bioreactor indicated that the Desulfovibrionales order, which have the ability to metabolize sulphate, was present. Within the transcriptome data, sulphur metabolism was confirmed to be present, involving gene products such as sulphite reductase (alpha and beta subunits), including the dissimilatory sulphite reductase alpha subunit (dsrA) and gamma subunit (dsrC), as well as the adenylylsulphate reductase (APSR) beta subunit. The bioreactor diversity also indicated that the Pseudomonadales, Rhizobiales, and Actinomycetales orders, which have the ability to fix nitrogen, were present. Transcriptome data confirmed that nitrogen metabolism was present, containing gene products such as the nitrate reductase catalytic subunit, the periplasmic nitrate reductase precursor, and 2-nitropropane dioxygenase. Chapter 7 244 For the methanogenic Archaea, the heterodisulphide reductase (HDR) which catalyses the reversible reduction of the heterodisulphide (CoM-S-S-CoB) of the methanogenic thiol-coenzymes such as coenzyme M (CoM-SH) and coenzyme B (CoB-SH) (Hedderich et al., 2005), were all present in the transcriptome, thus the survival of these groups could eventually have a possible effect on methane production. Several gene products involved in chemotaxis, regulation, and cell signaling were also detected such as the methyl-accepting chemotaxis protein (MCP), the aerotaxis receptor (Aer), and the signaling proteins CheY, CheZ, and CheV. Other gene products of interest found included the Fe/S oxidoreductases, involved in stress response and oxidative stress, the methyltransferases, Zn-dependent hydrolase/oxidoreductases, aldehyde ferredoxin oxidoreductases, Quinone oxidoreductases, ferredoxin, and GTPase – a putative metal chaperone involved in zinc homeostasis. Even though full pathways were not identified from the sequence analyses (Figure 7.17), several gene products were detected that are indicative of suspected active cycling in subsurface metabolisms, also found by Lau et al., (2014) and could allow the subsurface biome to deal with CCS conditions. Chapter 7 245 Figure 7.15: MG-RAST Krona diagram of the functional category hits distribution of the subsurface biome. Chapter 7 246 Figure 7.16: MG-RAST Krona diagram of CO2 fixation in the subsurface biome. Chapter 7 247 Figure 7.17: MG-RAST KeggMapper. The genes present within the subsurface biome are shown in the blue highlighted lines. Chapter 7 248 7.5 CONCLUSIONS According to the literature (Morozova et al., 2010; Basava-Reddi, 2012; Mu et al., 2014), the injection of sc-CO2 could cause a substantial decrease in the diversity in the subsurface, but ultimately the surviving microorganisms might be able to adapt and utilize the CO2. Using the continuous high pressure bioreactor, it was found that indeed a significant decrease in the diversity was observed however, several microorganisms were able to survive the CCS conditions likely due to their ability to form biofilms which protected them from these stressful conditions. It is known that microorganisms living in the extreme conditions of the deep subsurface environments display limited growth with exceptionally low metabolic activities and turnover times (Phelps et al., 1994; Teske, 2005; Jørgensen & D’Hondt, 2006; Jørgensen & Boetius, 2007; Onstott et al., 2009), therefore it was not unexpected that typical metabolic products were not observed during/after the six weeks of circulation under CCS conditions, even though these microorganisms remained viable. As expected, groups from the Bacteria and Archaea were able to survive the CCS conditions, but no Eukarya could be detected after six weeks. For the Archaea, only the Euryarchaeota phylum, consisting of the Methanobacteria and the Halobacteria classes, could be detected. The Bacteria remained the dominant domain, with the Proteobacteria still the main phylum present, similar to what has been found in the subsurface biome. The Firmicutes and the Bacteriodetes were also detected. In the acidic and high pressure environment created by the CCS conditions, metals, especially aluminium and iron, leached from the sandstone. This resulted in even more stressful conditions for the subsurface biome thus, restricting the already limited growth even further. This phenomenon should be considered before employing carbon sequestration, especially the influence it will have on the composition and cycling of the subsurface microbial communities. These extreme conditions limited successful RNA extractions and ultimately transcriptome analyses of the diversity, but data obtained for the subsurface biome indicated that gene products from sulphur metabolism, nitrogen metabolism, and carbon metabolism were present. The diversity within the bioreactor confirmed that microorganisms utilizing these metabolisms were present, and thus Chapter 7 249 specific cycling can be anticipated from the subsurface biome. 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The rise in population levels around the world has increased the energy demand and subsequent greenhouse gas emmissions. These activities are the main contributor to the increase in average temperatures worldwide, resulting in a global climate change problem. Therefore, carbon resources have to be managed more effectively in order to limit CO2 emissions. Current technologies include storing anthropogenic CO2 in geological formations. This might be a possible solution to mitigate the CO2 emissions on a scale that can stabilize the atmospheric greenhouse gases and make an impact on climate change. This study has illustrated that if these activities continue unmonitored, problems may occur as a result from the sequestered CO2. Studies have revealed that South Africa has the potential for storing CO2 in geological formations and that of the more than 400 million tonnes of annual carbon dioxide emissions in South Africa ~60% can be sequestered. However, it is well-established that the deep subsurface environment is a diverse and geologically complex domain, containing a considerable percentage of the Earth's biomass in the form of microorganisms. The deep terrestrial biosphere in South Africa is dominated by radiogenic noble gases and crustal-derived carbon sources, with microbial metabolisms dominated by methanogenesis, sulphate reduction, and fermentation. Therefore, the hydrological, microbiological, and geochemical processes of these subsurface microbial communities could result in natural chemical fluxes, influencing the quality of the water. Chapter 8 261 In order to survive these extreme environments with high pressures and temperatures, limited nutrient and energy sources, metal toxicity, as well as radioactive processes, the subsurface microbial communities had to evolve various mechanisms to adapt. In the process of adaptation, microorganisms can also change their surrounding environment in order to provide the necessary nutrients. As a result, the survival of microorganisms depends mainly on their ecological and metabolic capabilities. These subsurface microbial activities could potentially play a critical role in the cycling of carbon. Microbial metabolism under these conditions can substantially change the structure and chemical composition of rock formations, and potentially lead to changes in the porosity and permeability of the storage reservoir, resulting in leakage of the CO2, or even methane, since the CO2 may be fixed into organic matter by chemoautotrophs and converted into methane by the methanogens. This study evaluated subsurface microbial diversity from an environment of similar geological strata and depth as the sites proposed for carbon sequestration in South Africa. The biogeochemistry of the fissure water from the Star Diamonds mine in the Karoo sandstone was characterized over time. The fissure water was geochemically very similar over time, with a calculated conventional radiocarbon age of 24 320 (~131) years BP. Radiolysis of water could not be observed, possibly an indication that influx from older water sources are not relevant for this time period. Isotopic signatures revealed a possible mixing of thermogenic and biogenic gases, with positional and isotopic characteristics that could be consistent with production by methanogens utilizing the CO2 reduction pathway. The microbial diversity assessments confirmed that all three domains of life were present in the subsurface biome from the study site. The Bacterial domain was dominated by the Proteobacteria phylum, mainly consisted of Alpha-, Beta-, and Gammaproteobacteria classes but it also contained the Firmicutes, with the Bacilli and Clostridia classes. The Archaeal domain was dominated by the methanogens and halobacteria from the Euryarchaeota phylum, but also contained the Crenarchaeota. Chemolithoautotrophic microorganisms, as well as microorganisms with heterotrophic metabolisms were present in the subsurface biome and included the methanogens, which can fix CO2, sulphate-reducing bacteria, and microorganisms known to fix nitrogen. For the Eukaryal Chapter 8 262 domain, the main phyla present were Arthropoda, Streptophyta, Nematoda, Apicomplexa and Chlorophyta. When the subsurface biome was subjected to low pressure, in both nutrient rich and nutrient deprived environments, high biomass yields could be obtained for the nutrient rich conditions. However, for the nutrient deprived conditions, consisting of fissure water alone, exponential growth did not occur; the fissure water was not able to provide all the necessary nutrients for optimum growth. Changes in diversity can occur when subsurface biomes are exposed to different nutritional environments, as well as different physiological growth conditions. This was confirmed through diversity studies after the subsurface biome was grown under the various growth conditions. Although exponential growth did not occur under all the conditions tested, the microorganisms remained viable and metabolically active. Due to the extreme conditions of the deep subsurface environments, most of these microorganisms encounter energy limiting conditions, making them likely to be inactive or to display very low levels of metabolic activity. Microbial viability may ultimately prove to be a more important parameter than specific diversity changes or biomass yields for carbon cycling studies in the deep subsurface. To study the subsurface biome under conditions similar to what they could encounter during carbon sequestration, specifically designed bioreactors were used that were able to operate under higher pressures and temperatures. Under these conditions, the addition of CO2 resulted in it being in a supercritical state. Supercritical CO2 generally has harmful impacts on the majority of the microorganisms. However, throughout this research it was found that even though the microorganisms were not able to proliferate under these harsh conditions, they remained alive, even under 100 bar pressure, in the presence of 100% CO2 and in a nutrient deprived environment. This allowed for successful evaluations of the subsurface biome in an environment that mimics geological carbon sequestration. The presence of biofilm formations have been observed throughout these evaluations and could be the most important protection mechanisms used by microbial communities to survive the stress created by the presence of supercritical CO2. The presence of supercritical CO2 could cause a substantial decrease in the diversity of the deep subsurface, but ultimately the surviving microorganisms might be able to adapt Chapter 8 263 and utilize the CO2. To evaluate the influence of geological carbon sequestration on the subsurface microbial diversity and metabolic carbon cycling, a continuous high pressure bioreactor, specifically designed for this research, was used. These studies confirmed that a significant decrease in the diversity was indeed observed, but several microorganisms were able to survive these carbon sequestration conditions, most likely due to their ability to form biofilms. Even though several microorganisms remained viable, expected metabolic products were not observed during/after six weeks of circulation in the bioreactor, likely due to extremely low metabolic activities under carbon sequestration conditions. A substantial decrease in the diversity was observed after the six weeks of carbon sequestration conditions and as expected, groups from the Bacteria and Archaea were able to survive these conditions, but no Eukarya could be detected. The Bacteria remained the dominant domain, with the Proteobacteria the main phylum present, similar to what has been found in the subsurface biome. Firmicutes and Bacteriodetes were also detected. For the Archaea only the Euryarchaeota phylum, consisting of the Methanobacteria and the Halobacteria classes were observed. Transcriptome data obtained for the subsurface biome indicated that gene products from sulphur metabolism, nitrogen metabolism, and carbon metabolism were present and diversity analyses of the bioreactor confirmed that microorganisms, utilizing these metabolisms, were present. Thus, specific metabolic cycling can be expected from the subsurface biome, resulting in the production of formate, acetate, methane, and bicarbonates. The acidic and high pressure environment created by the carbon sequestration conditions allowed for significant leaching of metals from the sandstone, especially aluminium and iron. This increased the stress on the subsurface biome, limiting their already slow growth even further. This occurrence is of great importance and should be considered during carbon sequestration, since the influence it will have on the subsurface microbial communities could result in substantial impacts on the carbon cycling occurring in the deep subsurface. Ultimately, this project effectively demonstrated new considerations in terms of diversity, geological changes, and metabolic cycling that should also be considered during the planning and implementation of the carbon sequestration technology. Chapter 8 264 SUMMARY Climate change is a reality and is a consequence of too high levels of greenhouse gases being emitted into the atmosphere, especially CO2. This is happening mainly due to the burning of fossil fuels, but other human or industrial activities are also contributing. These additional emissions are responsible for the greenhouse effect, resulting in an increase in the global average temperature of the Earth's atmosphere. Industrialization and the rise in population levels around the world has increased the demand for energy and by meeting that demand more emissions are being released into the atmosphere and are responsible for an atmospheric increase of CO2 from 280 ppm around 1850 to 400 ppm today. This is creating long term, harmful conditions on the Earth, influencing the economy, the ecology, and human health. As an attempt to manage the carbon resources more efficiently, South Africa will be storing anthropogenic CO2 in geological formations. Within these deep subsurface environments, considered for carbon sequestration, highly diverse microbial communities can be found and the metabolic activities of these microorganisms are closely tied to the geochemical and mineralogical processes occurring within the subsurface. Therefore, to determine possible influences that the CO2 could have on the diversity and metabolic carbon cycling of these subsurface microorganisms, consideration was given to the interaction between the sequestered CO2 and the subsurface biome, as well as the geological environments. In order to possibly understand how these subsurface biomes will operate in situ, the biogeochemistry for a study site, similar to proposed sites for carbon sequestration in South Africa, was characterized comprehensively. The subsurface biome from the study site, used for this research, was found to contain microorganisms belonging to the Archaeal, the Bacterial, and the Eukaryal domains, but was dominated by the Bacteria, specifically the Proteobacteria. A specifically designed, continuous high pressure bioreactor was used to evaluate the influence of the stored CO2 on the subsurface microbial diversity and metabolic carbon cycling, by mimicking carbon sequestration conditions. Summary 265 Due to the extreme conditions of the deep subsurface, most of these microorganisms have to grow and survive in extreme energy limiting conditions, making them likely to be inactive or to display exceptionally low metabolic activities. Even though this research indicated that in most of the conditions tested the microorganisms were not able to multiply or to produce detectable amounts of expected metabolic products; they surprisingly remained viable. The most likely reason for this was due to the presence of biofilm formations which the microorganisms used as a possible protection mechanism towards resisting the harsh conditions they were subjected to. Ultimately, this indicated that viability might be more important, than specific diversity changes or biomass yields for carbon cycling studies in the deep subsurface. A considerable decrease in the diversity was observed after the biome was subjected to carbon sequestration conditions. Only groups from the Bacteria and Archaea were able to survive these conditions, but no Eukarya could be detected. Transcriptome data obtained for the subsurface biome indicated that gene products from sulphur metabolism, nitrogen metabolism, and carbon metabolism were present and diversity analyses of the bioreactor confirmed that microorganisms, utilizing these metabolisms, remained viable after six weeks under carbon sequestration conditions. These conditions, mainly due to the acidic environment created by the dissolved CO2, allowed for significant leaching of metals from the sandstone, especially aluminium and iron, creating more stressful conditions, thus restricting the growth of the subsurface biome even more. This could result in substantial impacts on the carbon cycling occurring in the deep subsurface, and should therefore be considered during carbon sequestration. Key words: climate change; carbon sequestration; geological storage; deep subsurface; metabolic carbon cycling; CO2 fixation; high pressure bioreactors; aluminium toxicity. Summary 266 OPSOMMING Klimaatsverandering is ‘n werklikheid en is die gevolg van te hoë vlakke van kweekhuisgasse, veral CO2, wat in die atmosfeer uitgestraal word. Dit gebeur hoofsaaklik as gevolg van die verbranding van fossielbrandstof, maar ander menslike en industriële aktiwiteite lewer ook ‘n bydrae. Hierdie addisionele uitstralings is verantwoordelik vir die kweekhuiseffek en dit veroorsaak gevolglik ‘n toename in die totale gemiddelde temperatuur van die aarde se atmosfeer. As gevolg van industrialisering en die die toename in die bevolkingsaanwas, het die vraag na energie verhoog en om in hierdie aanvraag te voorsien, word meer uitstralings in die atmosfeer gedoen, wat dus verantwoordelik is vir ‘n toename van atmosferiese CO2 van 280 dpm rondom 1850 tot 400 dpm vandag. Dit veroorsaak langtermyn skadelike toestande op die aarde wat weer ‘n invloed het op die ekonomie, die ekologie, en ook menslike gesondheid. Suid Afrika beplan om antropogenetiese CO2 in geologiese formasies te stoor, in ‘n poging om die koolstof voorrade meer doeltreffend te bestuur. Binne in hierdie diep ondergrondse omgewings, wat oorweeg word vir koolstofberging, word hoë mikrobiese bevolkings gevind en die metabolise aktiwiteite van hierdie mikroorganismes is baie nou verbind aan die geochemiese en mineralogiese prosesse wat in die ondergrondse omgewing plaasvind. Die interaksie tussen die opgeruimde CO2 en die ondergrondse bioom, asook die geologiese omgewings was ondersoek, om te bepaal wat die gevolglike invloed mag wees wat die CO2 op die mikrobiese diversiteit en metabolise koolstofsiklus mag hê. Om te probeer verstaan hoe hierdie ondergrondse bioom in situ funksioneer, was die biogeochemie vir ‘n studieterrein, soortgelyk aan die voorgestelde omgewings vir koolstofberging in Suid Afrika, volledig ondersoek. Dit was bevind dat die ondergrondse bioom van hierdie studieterrein mikroorganismes vanuit die Archaea, die Bakterieë, en die Eukariote bevat, maar was oorheers deur die Bakterieë, veral die Proteobakterieë. Koolstofbergingstoestande is nageboots deur die gebruik van ‘n spesiaal ontwerpte, deurlopende hoëdruk bioreaktor, om die invloed wat die gestoorde CO2 op die ondergrondse mikrobiese diversiteit en metabolise koolstofsiklus het, te evalueer. Opsomming 267 As gevolg van die uiterste toestande in die diep ondergrondse omgewing, moet die oorgrote meerderheid van hierdie mikroorganismes groei en oorleef in besondere energie beperkende toestande, wat veroorsaak dat hulle onaktief is, of besondere lae metaboliese aktiwiteite vertoon. Alhoewel hierdie studie aangedui het dat in die meeste van die toestande wat getoets is, die mikroorganismes nie in staat was om te vermeerder nie, of selfs waarneembare hoeveelhede van verwagte metabolise produkte te genereer nie, het hulle verbasend lewensvatbaar gebly. Die mees aanvaarbare verklaring hiervoor is dat die mikroorganismes biofilm gevorm het, wat hulle kon gebruik het as ‘n moontlike beskermingsmeganisme teen die uiterste toestande waaraan hulle blootgestel is. Dit dui uiteindelik daarop dat lewensvatbaarheid moontlik belangriker is as die spesifieke veranderinge in die diversiteit of die biomassa opbrengste, wanneer studies op die ondergrondse koolstofsiklus gedoen word. ‘n Merkbare afname in diversiteit was waarneembaar nadat die bioom blootgestel was aan koolstofbergingstoestande. Slegs groepe vanuit die Bakterieë en die Archaea was in staat om hierdie toestande te oorleef, terwyl geen Eukariote opgespoor kon word nie. Die transkriptoom data, wat verkry is vir die ondergrondse bioom, het aangedui dat geenprodukte van swaelmetabolisme, stikstofmetabolisme, en koolstofmetabolisme teenwoordig was en verskeie ontledings van die bioreaktor het bevestig dat mikroorganismes, wat gebruik maak van hierdie metabolismes, steeds lewensvatbaar was na ses weke onder koolstofbergingstoestande. Hierdie toestande, hoofsaaklik as gevolg van die suuromgewing wat deur die opgeloste CO2 geskep is, veroorsaak dat loging van metale uit die sandsteen, veral aluminium en yster, plaasvind, wat op hul beurt verdere stresvolle toestande veroorsaak en die groei van die ondergrondse bioom nog verder beperk. Dit kan ‘n baie groot invloed uitoefen op die ondergrondse koolstofsiklus en behoort gevolglik oorweeg te word tydens koolstofberging. Sleutelwoorde: klimaatsverandering; koolstofberging; geologiese berging; diep ondergrond; metabolise koolstofsiklus; CO2 fiksasie; hoëdruk bioreaktors; aluminium toksiteit. Opsomming 268